Regulation of Fimbrial Expression
IAN BLOMFIELD1 AND MARJAN VAN DER WOUDE2*
[SECTION EDITORS: ANDREAS J. BÄUMLER AND DAVID LOW]
Posted September 13, 2007
Department of Biosciences, University of Kent, Canterbury, Kent CT2 7NJ,1 and Department of Biology and the Hull York Medical School, IIU Area 12, University of York, P.O. Box 373, York YO10 5YW,2 United Kingdom
*Corresponding author. Mailing address: Department of Biology and the Hull York Medical School, IIU Area 12, University of York, P.O. Box 373, York YO10 5YW, United Kingdom. Phone: 44-(0)1904-328841. Fax: 44-(0)1904-328844. E-mail:
This e-mail address is being protected from spambots. You need JavaScript enabled to view it
.
Fimbriae, or pili, are proteinaceous structures extending from the surface of a bacterial cell that facilitate adhesion to abiotic and biotic surfaces. They are biochemically diverse, complex structures consisting of multiple proteins, including membrane anchors and major and minor subunits. Specific chaperones are often required for their assembly. The operons or gene clusters encoding these proteins frequently also encode regulatory proteins. The biochemistry of the fimbrial subunits determines the adhesive properties, and many specific receptors, along with the corresponding adhesin subunits, have been identified and well characterized. These studies, together with in vivo analyses of fimbrial mutants, indicate that most fimbriae are virulence factors that facilitate the colonization of specific host tissues, and some, including type 1 fimbriae of Escherichia coli, also facilitate adhesion to inorganic surfaces.
With few exceptions, fimbrial synthesis is encoded by a single operon. The metabolic drain on the bacterial cell of producing the abundant fimbriae is likely to be significant, and since neither E. coli nor Salmonella species are limited to the host environment for growth, regulation in response to signals indicative of the in vivo environment is to be expected. Furthermore, fimbria-mediated interaction of a bacterium with host cells can elicit both innate and adaptive antibacterial immune responses, and thus, even in a host, expression may not always be beneficial. For example, since fimbriae are antigenic, variable expression in a host may facilitate immune evasion. It is not surprising, therefore, to find that fimbrial production in E. coli and Salmonella enterica is under extensive environmental regulation. In this section, we review key features of the regulation of fimbrial gene expression in these two species.
Fimbrial gene expression, like most gene expression, can be controlled by transcriptional and posttranscriptional regulation. Transcriptional regulation includes classical regulatory mechanisms involving activators and repressors. Whether these regulators are global or local, and which environmental signals affect expression, varies per fimbrial operon, and most operons are controlled by multiple factors. Posttranscriptional regulation of fimbrial expression in the form of differential mRNA stability, which results in the differential expression of genes within a fimbrial operon, has been identified previously and is thought to ensure that the correct relative amounts of the various proteins are made. A recent study identified two small regulatory RNAs as regulators of some fimbrial operons, but the mechanism is not yet known (73). Other posttranscriptional mechanisms, for example, mechanisms involving RNA binding proteins, have not yet been identified.
Many fimbrial operons are also controlled by phase variation, and in the two species under consideration here, the phase variation mechanisms invariably affect transcription initiation. As a result of phase variation, expression is either in the off phase, when the operon is not expressed, or in the on phase, when the operon is expressed. The expression state is heritable but also reversible. The frequency of the switching event is characteristic of a specific operon and set of growth conditions and is between 1 in 10 cells per generation and 1 in 10,000 cells per generation. Thus, a mother cell that is in one phase (off or on) for the expression of a specific fimbrial type gives rise to progeny of which the majority remain off or on, yet a few will have switched to the opposite phase (Fig. 1). The switching events are random or stochastic in the sense that it is not possible to predict which individual cell will undergo a phase transition. In general, the frequency of the switch from on to off is higher than that of the switch in the reverse direction, suggesting that phase variation strictly limits the number of fimbriated cells present in the population. The phenotypic switch that results in phase variation from off to on is probably quite rapid, with fimbriae being formed within a generation. However, fimbriae are stable structures, and since there is little or no evidence of active shedding or retraction, a decrease in the number of existing fimbriae is presumably achieved only by dilution as cells divide. Thus, a switch from an on to an off phenotype may take several generations, lagging substantially behind the phase transition in the expression state.
The mechanisms underlying phase variation in different bacterial species are diverse and complex, but only two that control fimbrial phase variation in E. coli and Salmonella have been identified to date. These mechanisms are illustrated here by two paradigms: the fim and pap operons of E. coli. Notwithstanding these paradigms, the mechanisms determining the phase variation of several fimbrial operons in S. enterica serovars, such as lpf (106, 140), and in E. coli, such as the operon encoding 987P fimbriae, remain elusive and may prove to be novel.
The key difference between the mechanisms of fim and pap phase variation is that the regulation of fim is genetic since it involves a change in DNA sequence between on and off cells whereas the regulation of pap phase variation is epigenetic. Thus, pap phase variation does not involve changes in DNA sequence but is instead mediated by differences in DNA modifications at specific sites in the regulatory region. Both phase variation mechanisms are integrated with global regulatory systems that allow a response to environmental signals, either by leading to an altered switch frequency or by superimposing regulation on cells in the on phase. This regulation is discussed in further detail in the relevant sections.
The remainder of this chapter is divided into five sections, the first four of which are based on the key regulatory features; one fimbrial operon is used as a paradigm to illustrate each feature. Note that fimbrial operons are modular, as is evident by the absence of a strict relationship between the biochemical similarities of the subunits and the regulatory mechanisms. Thus, fimbriae that are discussed together in one section in this chapter based on shared regulatory principles may be unrelated based on the biochemical properties of the subunits. Furthermore, note that different isolates may contain genes and regulatory sequences that have diverged slightly, which can affect expression. The first section of this chapter describes the regulation of fimbrial operons controlled by AraC-like proteins, focusing on Rns. The second section describes the regulation of the csg operons encoding curli, the third section addresses the operons regulated by site-specific recombination as illustrated by a description of the fim operon, and the fourth section covers the pap-like family of operons regulated by epigenetic phase variation. For each paradigm, a comprehensive discussion presents all the regulatory features and factors affecting expression, and this discussion is followed by an overview of significant variations from the paradigm. In the last section of the chapter, an overview of recent insights into the regulatory networks that control the coordinated expression, temporally and possibly spatially, of fimbriae and flagella is presented.
The occurrence and distribution of fimbrial operons vary significantly among E. coli pathovars and even among the many Salmonella serovars. The paradigms that are discussed here are drawn mainly from E. coli because these have been studied extensively, and compared to the data on these paradigms, surprisingly little information on the regulation of the many Salmonella fimbrial operons is available. Details for the agf, fim (39, 69), and pef (136) operons in S. enterica are known, but differential regulation of many operons occurs. S. enterica serovar Typhimurium, for example, carries 13 putative fimbrial operons: agf (csg), fim, lpf, pef, bcf, stb, stc, std, stf, sth, sti, saf, and stj (93, 126). There is evidence of the expression of the following eight of these in vivo, but not under conditions tested in vitro: lpf, bcf, stb, stc, std, stf, sti, and sth. The three operons of which regulation has been studied previously (agf, fim, and pef) are expressed in vitro and also in vivo (92, 93). Thus, it is likely that regulatory mechanisms in addition to those described in this chapter exist, especially concerning signals for in vivo expression and for operons in which the regulatory sequence is not similar to those of well-studied systems.
Complementary information on fimbrial regulation can be found in other chapters, including those in section 8.3; specifically, information on curli is given in Chapter (Colonization of Abiotic Surfaces) and Chapter (Type 1 Fimbriae, Curli, and Antigen 43: Adhesion, Colonization, and Biofilm Formation) , data on K88, K99, and 987P fimbriae are presented in Chapter (Adhesins of Enterotoxigenic Escherichia coli Strains That Infect Animals), adhesins of diffusely adherent and enteroaggregative E. coli strains are discussed in Chapter (Adhesins of Diffusely Adherent and Enteroaggregative Escherichia coli), and information on adhesins of enterohemorrhagic E. coli is included in Chapter (Adhesins of Enterohemorrhagic Escherichia coli).
AraC-like transcriptional regulators activate the expression of some fimbriae (64), including CS1 and CS2 (regulated by Rns) (32), CS4 (regulated by CsvR) (47), 987P (regulated by FapR or FasH) (55, 107), colonization factor antigen I (regulated by CfaR) (33) in enterotoxigenic E. coli (ETEC), and aggregative adherence factors I and II (regulated by AggR) (135) in enteroaggregative E. coli. While the expression of many adhesins, including 978P fimbriae (195), is known to be controlled by phase variation, the expression of other adhesins regulated by AraC-like transcription factors does not appear to be under this control. The understanding of the regulation of CS1 by Rns is most advanced, and this system is described in more detail below.
Rns is a 26-kDa, plasmid-encoded protein required in trans for the expression of adhesins CS1 and CS2 (32). The rns gene is distinct from those required for the fimbrial structural components. Rns can substitute for CfaR and for FasH, and all of the AraC-like regulators listed above exhibit homology in their C-terminal regions to AraC, as do other members of the family (33, 130).
Rns activates its own expression from Prns, apparently to overcome the inhibitory effect of one or more negative regulatory elements that suppress rns expression (63). Rns requires both an upstream and a downstream binding site to activate its own promoter (132). Located 224.5 bp upstream of Prns, site 1 is farther away from the promoter than is typical for a bacterial transcriptional activator. In addition, the binding of Rns to a site centered at 43.5 bp downstream of the transcription start site is also required for the activation of rns transcription. This requirement for two binding sites may be a common feature of other AraC-like regulators that control virulence factors (123, 130). The activation of the CS1 structural genes by Rns, transcribed from Pcoo, appears to be more conventional, with binding sites centered 109.5 and 37.5 bp upstream of the promoter (131). DNase I footprint analysis of Pcoo reveals that Rns binds to degenerate, asymmetric recognition sequences, and DNA footprint analysis indicates that the protein interacts with two regions of the major groove along one face of the DNA, similar to other members of the AraC family. Rns may overcome the inhibitory effect of both H-NS and other local regulatory elements to activate Pcoo (133).
Although CS1 expression is affected by both aeration and temperature, a link between Rns and a response to these or other environmental signals has yet to be made (63, 177). However, Rns includes an amino-terminal domain of unknown function that may interact with the environment to regulate CS1 and CS2 expression.
Curli are adhesins that were initially identified by their ability to mediate the binding of Congo red and now are known to mediate binding to proteins like fibronectin (146) and to abiotic surfaces. In S. enterica serovars Typhimurium and Enteritidis, curli and cellulose expression is associated with the rdar morphotype, which reflects multicellular behavior. The term rdar stands for rough and dry colony morphology and the ability to bind Congo red, which are the originally identified features of this phenotype (162). In E. coli O157:H7 strain 4389R, a comparable but cellulose-negative morphotype was identified (190). In both E. coli and Salmonella, curli fimbriae are involved in biofilm formation (27, 69, 154, 155). The organization patterns of the genes required for the synthesis of curli fimbriae in E. coli and Salmonella are similar; in S. enterica serovars Typhimurium and Typhi, these genes have been designated csg and, especially in earlier reports, agf (for aggregative thin fimbriae), and in serovar Enteritidis, they are known as the SEF17-encoding genes (39, 164). The genes required for curli synthesis comprise two divergent operons, csgBA and csgDEFG (Fig. 2). The genes in the former operon encode the two structural proteins, and those in the latter code for a regulatory and a chaperone protein and two assembly proteins (76, 166). Specifically, the csgA gene encodes the major subunit, csgB encodes the nucleator, and csgD codes for a regulator. CsgG is an outer membrane lipoprotein that mediates the translocation of the subunits (160).
The regulation of curli expression appears to be in large part the same in E. coli and Salmonella and has been extensively studied in the context of biofilm formation. Multiple global regulators interact with the two csg promoters, resulting in a complex regulatory network that can fine tune the expression of curli in response to growth phase and numerous environmental stimuli. Known factors affecting curli production include increased expression due to nutrient starvation, low oxygen tension, alkaline conditions (pH 8.5), low osmolarity, and a low temperature (28°C) (reviewed in reference 70). However, most studies have examined regulation in only one of the two species, and the complete extent of variation is not known.
The csg intergenic regions of the two bacterial species have a relatively high level of sequence variation (164), likely contributing to species-specific regulation. For example, a role for the integration host factor (IHF) has been described only for Salmonella serovar Typhimurium (68, 69, 70, 165), and H-NS is a positive regulator in Salmonella serovar Typhimurium (69) whereas it is a negative regulator in E. coli (3, 145). Variable levels of curli expression among Salmonella serovar Typhimurium strains (166) and relatively high levels of curli expression in E coli O157:H7 (191) can be attributed to different alleles and specific mutations in the csg cluster. In contrast, the variable temperature dependence of curli expression, specifically, the absence of expression by E. coli K-12 at 37°C under non-biofilm-forming conditions (105, 146), can be attributed to a variation in the ompR allele (155, 201). Thus, significant variation in the expression of curli between the two species and even among isolates of the same species can occur.
The only regulatory protein encoded by the csg cluster is CsgD (AgfD), which has a LuxR- and FixJ-related helix-turn-helix DNA binding domain. CsgD was recently found to localize to the cytoplasmic membrane in E. coli (26), which is unusual for a transcriptional activator. The transcription start site for csgD in Salmonella serovar Typhimurium is 174 bp (164) and that in E. coli is 176 bp (76) upstream of the translation start site. The untranslated leader region is required for the maximal expression of a csgD-reporter fusion in Salmonella serovar Typhimurium and appears to be of particular relevance in the stationary phase (68).
The regulation of curli expression can occur at either or both csg promoters, and thus, environmental control of the expression of the subunits can be mediated at either promoter. CsgD is a positive regulator of csgBA transcription (27, 76, 154) and of its own promoter (PcsgD). Furthermore, CsgD is unusual for a fimbrial operon-encoded regulator in that it participates in both positive and negative regulation of as many as 24 genes outside the csg operon in E. coli. These include several genes involved in biofilm formation, like fecR, which affects aggregation, and yoaD, which controls cellulose production and also affects aggregation (26, 27). Furthermore, CsgD affects glyA expression and thus affects glycine biosynthesis (35), which may be related to the high glycine content of curli fibers (35). In Salmonella, CsgD also participates in the regulation of genes that are involved in biofilm formation in addition to the csg genes, including those for cellulose biosynthesis in Salmonella serovar Typhimurium (166, 218) and the outer membrane protein BapA in Salmonella serovar Enteritidis (113).
Three two-component regulatory systems are known to affect transcription at one or both of the csg promoters. The EnvZ-OmpR system (reference 201; also see Chapter Colonization of Abiotic Surfaces) is required in both E. coli and Salmonella for csgD transcriptional activation, and consistent with this requirement, there is a promoter-proximal site for OmpR-P (Fig. 2) (69, 98, 154, 164). In Salmonella serovar Typhimurium, this D1 site is necessary and sufficient for activation (68). Five additional OmpR-P binding sites in the intergenic region in Salmonella serovar Typhimurium contribute to transcriptional repression specifically in response to high oxygen levels, (69, 70). A seventh binding site D7 at 406 to 429 upstream of the csgD transcription start site was also identified and contributes to repression under microaerophilic conidtions (68). IHF competes with OmpR-P for binding sites 3 through 6 under conditions of low oxygen tension and, hence, apparently antagonizes negative control by OmpR-P (69). The molecular basis of the link between OmpR-P-dependent regulation and oxygen tension remains to be identified and may involve variations in the level of OmpR-P. However, the posttranscriptional control of csg in response to OmpR-P may also have to be considered. Specifically, a microarray study determined that the levels of expression of all csg genes in E. coli decrease in response to increased expression of the omrA and omrB (formerly gyrA and gyrB) small regulatory RNAs. OmpR-P was previously shown to increase the expression of these regulatory RNAs (73), and thus, some OmpR-dependent regulation of csg may be indirect.
The OmpR-P binding site in E. coli overlaps one of the binding sites of the second two-component regulatory system involved in curli regulation, CpxAR (50, 98, 154), which senses envelope stress. The Cpx system mediates the repression of the csgD promoter (199), and the positions of the binding sites are consistent with this finding (98). A deletion of the tol genes that are involved in membrane integrity results in fewer curli and the repression of the transcription of csgBA, presumably as a result of envelope stress. This regulation requires repression mediated by the third two-component regulatory system, which is the Rcs system (60, 199). However, in the tol mutant, csgD transcription increases, and this effect requires OmpR (199). Since this increase in transcription does not result in an increase in curli production, the relevance of the upregulation of csgD transcription in this mutant may relate to its role in regulating other operons. A microarray analysis showed that negative regulation of csgD occurs upon the activation of the RcsC sensor kinase. Subsequently, a study using a reporter gene demonstrated that both RcsA and RcsB are moderate repressors of the csgD promoter and strong repressors of the csgB promoter (199). Interestingly, RcsC is activated on solid surfaces, and this activation may result in the downregulation of curli production (60). How this finding relates to the role of curli in biofilm formation remains to be determined.
Curli production in E. coli is repressed in response to high osmolarity (154), and this response can be attributed to the repression of promoter activity at csgD as indicated by analyses with reporter fusions. Even though the EnvZ-OmpR two-component pathway mediates the response to osmolarity and OmpR-P activates transcription at PcsgD (154), the regulation of curli in response to changes in osmolarity does not appear to involve OmpR-P. Instead, the only regulators in E. coli identified to date are H-NS and CpxR-P (98). In medium with a high sucrose content, the downregulation of csgD is independent of the Cpx system and is mediated through H-NS (98). In contrast, in high-salt-content medium, repression requires CpxR (98), which may indicate that CpxR mediates a response to ionic strength rather than osmolarity. In Salmonella serovar Typhimurium, csgD is downregulated by high salt concentrations but not by high sucrose levels (164), also suggesting that regulation is a result of the sensing of the ionic strength rather than osmolarity.
A recent report indicates that curli production in E. coli is repressed in response to the increase in the intracellular concentration of N-acetylglucosamine-6-phosphate (GlcNAc-6P) that results from a mutation in nagA or in response to exogenous GlcNAc (9). This finding was confirmed by measuring CsgA and CsgD protein levels, as well as the levels of expression of reporter gene transcriptional fusions. Interestingly, GlcNAc, via its ability to inactivate NagC, also suppresses type 1 fimbriae (181) (see also the section on fim regulation in this chapter, below). However, several lines of evidence suggest that NagC does not mediate the effect of GlcNAc-6P on curli production, and the molecular mechanism that results in the transcriptional repression of curli remains to be determined (9).
Curli production in both E. coli and Salmonella serovar Typhimurium increases as nutrients are depleted and, thus, increases in late exponential and stationary growth phases. In the stationary phase, the transcription of both csg promoters in E. coli (76) and in Salmonella serovar Typhimurium (164) involves sigma factor RpoS. A previous mutational analysis showed that, in E. coli, the gene crl is required for curli production (4). It was initially thought that crl encoded the curli subunit, hence the name Crl for the gene product (146). However, it is now clear that Crl is a regulatory factor that enhances the expression of a subset of the RpoS-dependent genes by directly interacting with RpoS and that remains associated in the RNA polymerase RpoS holoenzyme (Eσs) (21, 153). Later studies of Salmonella serovar Typhimurium using reporter fusion analyses showed that Crl is required for the maximal expression of both csgD and csgB (159). In vitro transcription analyses showed that Crl increases transcription at csgD via Eσs, and that Crl mediates this activation at the csgD promoter by enhancing open complex formation (159). In vitro transcription of the csgB promoter has not been successful, suggesting that an additional regulatory factor needs to be identified. Therefore, it is not clear if activation at the csgB promoter is indirect or direct (21, 159). However, in vitro, Crl affects the DNase I footprint and the recruitment of Eσs at the csgB promoter (21). The expression of crl increases at lower temperatures and in stationary phase (159), and this increase appears to contribute to the temperature- and growth phase-dependent expression of curli (21). Temperature-dependent regulation by crl may be in addition to temperature-dependent effects on RpoS levels mediated through the DsrA small RNA (sRNA), especially in E. coli (reviewed in references 96 and 157). This crl dependence of curli expression is, however, a trait that varies among isolates. For example, in an avian E. coli isolate, curli production is not affected by a crl mutation (156). In most Shigella species isolates, apparent pathoadaptive mutations in crl result in the absence of curli expression (170).
The cyclic dinucleotide 3',5'-cyclic di-GMP (c-di-GMP) has been recently identified as a global second messenger in bacteria (163), and regulation by c-di-GMP levels underlies the rdar morphotype of Salmonella serovar Typhimurium (99). Even though the mechanism of c-di-GMP-dependent regulation in general is unclear, this regulation of the rdar morphotype is at least in part due to the coordinated regulation of cellulose and curli production. This link was discovered by analyzing the role of GGDEF-EAL domain proteins, which are involved in c-di-GMP turnover, in the rdar morphotype. Specifically, the overexpression of AdrA, which results in increased levels of this second messenger, led to enhanced curli expression (99). This enhancement correlates with increased levels of both CsgA and CsgD proteins (99), which correlate with increases in transcription initiation at the csgBA promoter and levels of csgD transcripts. CsgD protein levels, however, also diminish in isolates with lower levels of c-di-GMP, suggesting that posttranscriptional regulation may also occur. The contribution of individual GGDEF-EAL domain proteins to regulation appears to be complex, since individual mutations have different effects on CsgD and CsgA levels. For example, an adrA mutation results in a decreased level of CsgA but does not affect the level of CsgD (99). Interestingly, CsgD activates the expression of AdrA (218), suggesting that under certain conditions a positive feedback loop may be established. It seems likely that this feedback loop is an important regulatory feature coordinating the expression of adhesins in bacteria.
Despite a significant increase in our understanding of curli regulation, it is likely that additional regulatory factors and signals will be identified. For example, a mutation in the conserved yehV gene, designated mlrA, inhibits the transcription of csgD and csgBA in Salmonella serovar Typhimurium, but it is not known to which environmental signals this inhibition mediates a response (28). When these findings are taken together, it is clear that curli production is controlled by transcriptional and posttranscriptional mechanisms and is under the control of a multitude of regulatory signals and factors. Since curli are important in biofilm formation, this complexity probably reflects the general principle that the expression of factors involved in this multicellular behavior is tightly controlled.
The regulation of curli production is also discussed in detail in Chapter (Colonization of Abiotic Surfaces), which includes an additional overview of environmental signals that affect curli expression and a detailed explanation of the mode of action of OmpR. In Chapter (Type 1 Fimbriae, Curli, and Antigen 43: Adhesion, Colonization, and Biofilm Formation), regulation in the context of adhesion to abiotic surfaces and biofilm formation is reviewed. Curli expression specifically in enterohemorrhagic E. coli O157:H7 is discussed in Chapter (Adhesins of Enterohemorrhagic Escherichia coli), and curli expression in the context of glycine biosynthesis is discussed in Chapter (Regulation of Serine, Glycine, and One-Carbon Biosynthesis).
The morphology of type 1 (SEF21) fimbriae of Salmonella serovars Typhimurium and Enteritidis resembles that of the E. coli type 1 fimbriae, and like the latter, SEF21 fimbriae mediate mannose-sensitive hemagglutination (129). However, the arrangement of the Salmonella fim gene cluster and the regulatory mechanism are quite different from those of the E. coli fim operon. The Salmonella serovar Typhimurium fim cluster contains six genes in an operon, fimAICDHF, of which fimA encodes the main fimbrial subunit. However, fim expression is regulated by three genes downstream of, and organized as a convergent cluster with, fimA to fimF, fimW, fimY, fimZ, and a fourth gene, fimU, which is divergent from fimW. FimZ, FimY, and fimU are required for type 1 fimbrial synthesis (187, 188, 216), whereas FimW is a negative regulator (189). The expression of these type 1 fimbriae appears to phase vary since a clonal population can be heterogenous for expression, but a mechanism remains to be identified (36).
FimZ has homology to the DNA binding proteins or response regulators of two-component regulatory systems, with a predicted helix-turn-helix DNA binding motif and a response-regulatory motif. FimZ is required for the expression of the major fimbrial subunit gene fimA (216) and for the expression of fimY (187). In a DNase I footprint assay, FimZ bound to fimA regulatory-region DNA in vitro (216), protecting the region from –47 to –98 relative to the transcription start site, which is a binding region similar to those found in other classical activators (217). FimZ also enhances its own expression, as determined in vivo by using a reporter fusion. However, no binding of FimZ to its own regulatory region could be detected, and the mechanism underlying this autoregulation remains uncertain (217). There is no adjacent kinase, and whether or not phosphorylation plays a role in FimZ activity like it does in the activity of other response regulator proteins is not known. The expression of fimA and fimZ also requires the second activator in the cluster, FimY (187, 216, 217). The amino acid sequence has, however, only limited homology to those of DNA binding proteins (187). Thus, FimZ and FimY are both required for fim expression and appear to be coactivators. However, no DNA binding activity could be detected in vitro with FimY and fim promoters, and the mechanism of activation remains to be determined (187).
The third regulatory protein, FimW, also has limited homology to prokaryotic regulators and functions as a negative regulator of fim expression. No binding to any of the fim promoter regions could be detected in vitro, but in a previous study using a two-hybrid system, interaction between FimW and the activator FimZ was shown (189). Therefore, FimW may exert negative regulation by interfering with FimZ activation or DNA binding. Interestingly, FimW expression increases during growth on solid medium and therefore may be involved in the observed downregulation of fim expression under these growth conditions (189).
The fourth regulatory gene, fimU, encodes a rare arginine tRNA, tRNAArg(UCU). Five rare arginine AGA codons are within fim, of which three reside at the 5' end of fimY. Indeed, fimU is required for the translation of a FimY fusion protein, and type 1 fimbriae are not expressed by a fimU mutant (186, 188). The fimW gene also contains two rare arginine AGA codons, but translation is not affected by a fimU mutation. The fimU dependency may link type 1 expression to the nutritional status of the cell. A mutation in fimU in Salmonella serovar Enteritidis affects the expression of two fimbrial operons, specifically, those of type 1 (SEF21) and SEF14 fimbriae (38).
Like the majority of fimbrial operons in E. coli, type 1 fimbriation is controlled by phase variation (25). Phase switching involves the reversible inversion of a short (~314-bp) element of DNA (the fim switch) by site-specific recombination (2). While this system in E. coli had appeared to be restricted to type 1 fimbriation, CS18 fimbriae of enteropathogenic E. coli are now known to be controlled in a comparable way (88).
The fim switch contains a promoter directed towards the fimbrial subunit, usher, and chaperone genes in the on, but not in the off, orientation (56, 147). Two recombinases, FimB and FimE, catalyze the inversion reactions (66, 108), albeit with different specificities and at different frequencies within the cell (17, 30, 65, 66, 108, 110, 124, 125, 149, 176, 185). The expression of fimE is, like that of the fimbrial subunits, controlled by the fim switch (83, 97, 178, 182). Thus, although the invertible element lies downstream of fimE (Fig. 3), it nevertheless suppresses fimE expression in fimbrial-off-phase cells. Site-specific DNA binding and bending proteins IHF and Lrp are auxiliary factors in the recombination reactions (15, 16, 52, 57, 67, 111, 161), and the inversion is affected additionally both by H-NS (48, 49, 82, 104, 144, 148, 184) and by transcription in the vicinity of the invertible element (144). Phase switching is regulated by environmental signals and is coordinated with Pap (86, 87, 213). The responses of the fim switch to the amino acids leucine and alanine and to amino sugars N-acetylneuraminic acid (Neu5Ac) and GlcNAc have been studied in the most detail (58, 65, 67, 111, 161, 181), but other factors, such as temperature, osmolarity, and pH, also control the system (65, 173). In addition, off-to-on phase switching has recently been shown to be stimulated by the alarmones ppGpp and pppGpp (1).
The region upstream of fim contains a large (1.4-kbp) intergenic region (Fig. 3), yet the expression of fimB, the first gene in the cluster, is coordinated with that of the divergently transcribed nanC (formerly yjhA) gene (40, 181). Shared cis-acting regulatory elements that overlap the nanC promoter bind Neu5Ac- and GlcNAc-6P-responsive proteins NanR and NagC, respectively. The binding of NanR and NagC to these regulatory elements is alternate, as indicated by methylation protection of Dam sequences in these binding regions (58, 181).
In comparison to the regulation of the fim switch, little is known about factors which control fim expression in cells in the on phase. However, when the invertible element is locked in the on orientation by mutation or when it is replaced altogether with the Ptac promoter, the resulting strains still alternate between fimbriated and afimbriated phases (124). Thus, the fim switch in the on orientation is necessary, but not sufficient, for fimbrial biosynthesis, and a posttranscriptional regulatory mechanism apparently also controls the phase variation of the adhesin. The phase variation of type 1 fimbriation is hence determined at multiple levels and, rather than occurring at a single fixed frequency, is regulated by multiple environmental signals. Moreover, to make matters more complicated, the regulation of fim phase variation in uropathogenic E. coli (UPEC) strains shows considerable strain-to-strain variation (91, 114, 117, 175). Such variation can arise via sequence heterogeneity at fim (114), but cross talk between fimbrial operons (213), as well as the presence of additional recombinases in some strains (29, 215), presumably also contributes to the effect.
fim
Transcription Units and the Invertible Element.
The fim genes, located on the E. coli K-12 genome at centisome 97.83 to 98.00, are transcribed in a clockwise direction (62) (Fig. 3). The invertible element is bound by two 9-bp inverted repeats (termed inverted repeat left [IRL] and inverted repeat right [IRR]) of 5'-TTGGCCCCA that flank a 296-bp region in E. coli K-12 (2). fimB and fimE, which encode the two fim recombinases, are transcribed separately from each other, and the fim invertible element (fimS) lies downstream of fimE (Fig. 3). In on-phase cells, the fimE transcript reads through fimS, whereas it is thought to terminate within the invertible element in the off phase (83, 97). Notwithstanding this pattern, the principle promoter for the fimbrial structural and assembly genes, termed PfimA, lies within the invertible element (147) (Fig. 4). PfimA overlaps the recombinase binding sites at IRR in the on orientation, and recombinase binding appears to inhibit transcription initiation (53, 176). Furthermore, differences in the nucleotide sequences of 5' ends of transcripts originating from this promoter in on and off cells may affect the stability of the RNA produced (147).
Site-Specific Recombination Controlling fim Phase Variation. (i) Recombinase Proteins and Their Activity.
FimB and FimE are small (around 23-kDa) basic proteins that show around 48% amino acid identity (108). The proteins bind, albeit with different affinities, to sites that flank, but only partially overlap, IRR and IRL (66). They each contain the conserved tetrad of amino acids (Arg-47 and Arg-41, His-141 and His-136, Arg-144 and Arg-139, and Tyr-176 and Tyr-171 for FimB and FimE, respectively) essential for catalysis by lambda integrase and related recombinases (176).
The binding sites for FimB and FimE adhere closely to the consensus sequence 5'-ANNAGACANTTNGG, and the central 5'-CA sequence is conserved fully (66). Like for many other recombinases, mutations that destroy the DNA homology within the region of strand cleavage and exchange (uncharacterized for IRL and IRR of fimS) prevent recombination and, hence, lock fimS either on or off (66, 124).
Although FimB and FimE catalyze the inversion of fimS in either direction in E. coli K-12 strain MG1655, inversion from on to off occurs at a much higher frequency than inversion in the opposite direction (around 0.3 and 0.001 times per cell per generation, respectively, in aerated MOPS [morpholinepropanesulfonic acid]-rich defined medium at 37°C). FimE is required for rapid on-to-off inversion and has a marked preference for fimS in the on orientation (17, 65, 108, 124, 125, 149). In contrast, FimB promotes inversion at lower frequencies and usually with little directional bias. FimB recombination, which can vary over several orders of magnitude under different conditions, thus determines the proportion of fimbriated bacteria within the population of strain MG1655 cells.
In addition to FimB and FimE, many uropathogenic strains encode proteins that show significant homology to the fim recombinases (29). IpuA (48% identity to FimB), IpuB (49% identity to FimB), and IpbA (55% identity to FimB) are all encoded by UPEC strain CFT073 (29, 202), and both IpbA and IpuA promote the inversion of fimS from off to on when supplied in trans on a multicopy plasmid (29). Of particular interest, IpuA permits the off-to-on inversion of fimS in a mutant of CFT073 that lacks all of the other recombinases both in vitro and in a mouse model of urinary tract infection (29). lpuA is found in a significant number of UPEC strains (36%) but is generally absent from commensals, so LpuA in addition to FimB may control type 1 fimbrial phase variation in many urinary tract infections. IpbA (also known as HbiF) has also been identified previously in a pathogenic E. coli K1 strain capable of causing meningitis in neonatal rats (215), and when expressed as a result of mutation, the protein is also able to catalyze the inversion of fimS in this background.
(ii) FimE Specificity and Orientational Control of fimE Expression.
In a fimB mutant background, fimS appears to be usually locked in the off orientation (17, 125, 185). An analysis of this phenomenon reveals not only that FimE has a preference for fimS in the on orientation as a substrate for recombination but also that fimE expression is inhibited in off-phase cells (83, 97, 110, 178, 182). In the absence of fimE, rates of on-to-off inversion are lowered around 100-fold, and thus, FimE dominates phase switching from the fimbriated to the afimbriated phase (17, 65, 124).
With a plasmid substrate in which fimS is flanked only by the external binding sites for FimE and with the recombinase supplied in trans, a strong bias for recombination in the on-to-off direction is still observed (110). Furthermore, an exchange of either the internal or external binding sites for the recombinase results in a reversal of specificity in this system. However, when additional fim sequences that flank the external recombinase binding sites are included, some loss of specificity is observed when the external binding sites are exchanged. FimE specificity is thus dependent on sequence differences between the recombinase binding sites but is also moderated further by additional sequences outside of fimS that remain to be identified.
FimE binds with differential affinities to IRL and IRR depending upon the orientation of fimS, with IRL in the off orientation having the lowest affinity (66). Thus, one simple explanation for why FimE is unable to carry out off-to-on recombination effectively is that the recombinase fails to bind to IRL in the off orientation at normal physiological concentrations. However, a mutation in FimE, R59K, that produces a loss in recombinase specificity does not alter the protein’s DNA binding affinity (176). Therefore, differential binding of the protein to fimS in the on and off orientations seems unable to account fully for FimE specificity.
fimS lies downstream of fimE, with the open reading frame for the recombinase gene ending just 51 bp from IRL. When a recombinase binding site mutation that reverses FimE specificity for the plasmid substrates described above is substituted for wild-type fimS in the chromosome, rather than an alteration in specificity as expected, no recombination is detected (110). This effect occurs because fimE expression is inhibited by the invertible element when it is both in the off orientation and in cis to the recombinase gene (89). fimE mRNA continues into fimS (182), and Rho-dependent transcription termination apparently produces a 3' end that allows rapid and selective degradation of fimE mRNA in off-phase cells (83, 97).
Under conditions that are selective for type 1 fimbriae, a low level of FimE-catalyzed off-to-on recombination in the absence of FimB can be detected (185). However, although FimE may contribute to off-to-on phase variation, as detailed above, several mechanisms exist to prevent FimE from catalyzing this inversion. The dual recombinase system that controls fim seems thus to have evolved to allow high rates of on-to-off phase switching without compromising tight control over much lower levels of switching in the opposite direction.
(iii) H-NS, FimB Specificity, and Thermoregulation.
Mutations that inactivate hns increase the frequency of FimB recombination substantially (82, 104, 184). H-NS binds to the fimB promoter region to repress fimB transcription but does not affect FimB recombination in cell extracts (49, 147, 148). At least part of the effect of H-NS on FimB recombination is hence explicable by an indirect effect of the protein on fimB expression. Notwithstanding this effect, hns mutations that disrupt the protein’s DNA binding activity fail to enhance FimB recombination fully even though they do increase fimB expression (48). Thus, H-NS may also bind to another protein to form a complex that inhibits FimB recombination more directly (48).
Transcription from a promoter inserted within fimE inhibits FimB recombination specifically from off to on (144). The effect observed requires PfimA, as well as H-NS. Since the overexpression of fimB suppresses the directional bias, the effect of H-NS seems likely to be indirect. While the significance of these observations is unclear, mutations in an IHF site situated between fimE and IRL (IHF site 1; see below) can also affect FimB specificity, and novobiocin, an inhibitor of the supercoiling activity of DNA gyrase, imparts a directional bias on FimB recombination in favor of off-to-on switching (53, 54, 114). FimB recombination is affected by mutations in topA, and DNA supercoiling in the vicinity of fimS may affect FimB recombination (54, 144). Although the effect of H-NS on FimE recombination has been investigated in less detail, the protein also represses fimE expression and inhibits rapid on-to-off inversion (148).
Like pap, fim phase variation is regulated in response to temperature (67). H-NS is reported to affect fimB and fimE expression differentially depending on the temperature, and this effect may explain how the fim recombinases are thermoregulated (148). Whereas FimE recombination decreases as temperature increases between 28 and 42°C, FimB recombination is optimal at around 37°C (67). However, in another study, it was reported that the level of FimB recombination at 30°C is higher than that at 37°C, indicating that strain variation or other factors affect the thermoregulation of FimB recombination (51).
(iv) IHF and Lrp and Regulation of the fim Switch in Response to the Branched-Chain Amino Acids and Alanine.
For strand exchange between IRL and IRR to occur, the alignment of the two inverted repeats requires substantial DNA bending. Global regulators IHF and Lrp bind to elements within the invertible region, and the sharp (>180°), in-phase bends that they produce in combination are thought to facilitate inversion. As expected, mutations in either ihf alpha or ihf beta or in lrp diminish both FimB and FimE recombination, as do those in the single IHF binding site (IHF site 2) that is situated closer to IRR than to IRL in the on orientation (15, 16, 52, 57, 67, 111, 161) (Fig. 4). Lrp binds cooperatively to three sites (Lrp sites 1 to 3) within the invertible element, situated closer to IRL in the on orientation (161). As described below, the occupancy of Lrp site 3 regulates both FimB and FimE recombination.
As noted above, IHF also binds to a site (IHF site 1) between fimE and IRL. Mutations in this element have a greater effect on FimB than FimE recombination (16), and IHF bound to site 1 may stimulate FimB-catalyzed off-to-on inversion preferentially (53, 114).
Leucine and alanine and, to a lesser extent, isoleucine and valine stimulate both FimB and FimE recombination (65). Leucine and alanine selectively inhibit Lrp binding to site 3 in vitro, promoting a nucleoprotein complex in which the protein remains bound to sites 1 and 2 (complex 2) at the expense of a complex in which all three sites are occupied (complex 1) (161). Mutations that diminish Lrp binding to site 1 or 2 inhibit recombination, irrespective of the presence of the amino acids. In contrast, those that produce a loss of site 3 binding stimulate inversion but do so selectively in medium lacking the amino acids. Furthermore, a mutation that increases Lrp binding to site 3 inhibits recombination in the absence of the amino acids and diminishes their stimulatory effect (111). Thus, the branched-chain amino acids and alanine stimulate fim phase variation by favoring the formation of an alternative Lrp-DNA complex (complex 2) that enhances recombination. Although it is unclear why Lrp complex 2 enhances recombination more than complex 1, further analysis revealed that the nucleotide sequence between Lrp site 3 and the IHF binding site contains additional elements that regulate the amino acid response (111). The response to the amino acids may thus be controlled by environmental or physiological cues that remain to be identified.
fimB and fimE each contain an unusually high proportion of poorly translated codons, indicating that the gene products are likely to be expressed at low levels (158). The replacement of the UUG (LeuX-decoded) codons of fimB with the more frequently used LeuV-decoded codon, CUG, does indeed increase fimB expression, as is evident through increased FimB-dependent recombination. Furthermore, under some growth conditions, mutations in lrp increase fimB expression (15). Although the significance of these observations is unclear, it seems probable that low levels of leucine inhibit off-to-on phase variation by limiting fimB expression, as well as by altering the interaction of Lrp with site 3 in fimS.
(v) Regulation of fimB Expression by the Alarmones ppGpp and pppGpp.
Mutants unable to synthesize (p)ppGpp show decreased expression of type 1 fimbriae, and this effect is explicable by a corresponding deficit in fimB transcription (1). Several investigators have reported previously that fimB transcription initiates from two promoters (1, 49, 147, 174), but only one of these (P2) is affected by (p)ppGpp (1). The response to (p)ppGpp is independent of DskA, a factor implicated in the regulation of many promoters by the alarmones (150).
Starvation for amino acids induces the stringent response and, hence, (p)ppGpp production. Since lrp is also stimulated by (p)ppGpp (112), starvation for the branched-chain amino acids likely induces opposing effects on fimB expression. Moreover, higher levels of Lrp should inhibit FimB recombination directly (111), and further work is needed to clarify the role of (p)ppGpp in regulating off-to-on phase variation under different environmental conditions.
(vi) NanR and NagC and Regulation of fimB Expression by Neu5Ac and GlcNAc.
fimB is separated from the divergently transcribed nanC gene (formally known as yjhA) by one of the largest (1.4-kbp) intergenic regions in E. coli (40) (Fig. 3 and Fig. 5). Despite this organization, a deletion of 252 bp (termed Δ2) more than 600 bp upstream of the fimB open reading frame decreases FimB recombination more than 18-fold (58). The effect is seen only when the region deleted is in cis to fimB, and it is associated with a fivefold decrease in fimB expression.
A mutational analysis of the sequences within Δ2 located two specific regions (originally termed region 1 and region 2 or NagC1 but more recently renamed ONR and ONC1) that stimulate fimB expression (58). Mutations in ONR have a lesser effect than those in ONC1, and a combination of mutations in both regions decreases fimB expression the most. ONR (underlined) falls within a 27-bp sequence (5'-ACCTTTATACCTGTTATACCAGATCAA) conserved upstream of both the nan and yjhBC operons. One of the homologs of ONR overlaps the nan operon promoter, and both this element and ONR bind the sialic acid-responsive regulator NanR (100, 181). As expected, both fimB expression and FimB recombination are inhibited by Neu5Ac (58).
ONR encompasses, and ONC1 lies immediately next to, Dam methylation sites (termed GATCNanR and GATCNagC, respectively) that are protected from modification in a subpopulation of cells (58, 181). The pattern of protection and methylation indicates that the simultaneous protection of both sites occurs rarely, if at all. Whereas a mutation of nanR produces a loss of methylation protection at GATCNanR specifically, Neu5Ac inhibits protection at both GATCNanR and GATCNagC. Further analysis shows that ONC1 binds the GlcNAc-6P-responsive protein NagC and that exogenous GlcNAc inhibits fimB expression. As expected, NagC is required for methylation protection at GATCNagC, and the addition of GlcNAc, which generates GlcNAc-6P upon uptake, to medium inhibits the methylation protection of GATCNagC but not GATCNanR. Since Neu5Ac catabolism generates GlcNAc-6P, the initially perplexing observation that Neu5Ac inhibits methylation protection at both sites is readily explained. Taken together, these observations support a model in which fimB expression is activated alternately by NanR and NagC, with GlcNAc inhibiting one, and Neu5Ac inhibiting both, of these pathways (181).
NagC sites characteristically function in pairs (152), but although NagC binds to a second element (ONC2; originally termed NagC2) situated 212 bp closer to fimB than ONC1, the cooperative binding of NagC to ONC1 and ONC2 was not apparent in vitro in previous studies, and a 304-bp deletion that included ONC2 (Δ3) did not diminish fimB expression (57, 178). However, further work showed that a point mutation in ONC2 both diminishes fimB expression and results in a loss of the methylation protection of GATCNagC (183). These apparent discrepancies were resolved by showing that NanR compensates for the loss of NagC activation of fimB in the Δ3 deletion mutant background. Furthermore, IHF binds to a site (ibs) midway between ONC1 and ONC2 to facilitate the cooperative binding of NagC to its two operator sites. IHF thus controls fimB expression and, in addition, the recombinase activity of both FimB and FimE. How NagC-IHF and NanR are able to activate fimB expression from distant activator sites situated far upstream of the fimB promoter remains to be determined.
Switching between NanR and NagC binding is rapid, and it has thus far not been possible to isolate cultures enriched with cells in which one or the other regulator alone is bound. Nevertheless, if NanR activates fimB expression to a lesser degree than NagC, as it appears to, levels of fimB expression presumably fluctuate as binding switches from one protein to the other.
While ONR coincides with the −10 region of a promoter for the divergently transcribed nanC gene, ONC1 encompasses a Crp binding site thought to activate nanC expression (40). The expression of fimB and nanC are thus coordinately controlled via a common regulatory region, with both NanR and NagC repressing nanC. NanC is a sialic acid-specific porin, and while fimA expression is suppressed by the OmrA and OmrB regulatory RNAs, that of nanA is enhanced by these factors (73). Although why type 1 fimbriation and Neu5Ac metabolism are coregulated also remains to be determined, coordination is apparently achieved at multiple levels.
Control of fim Expression in Cells at the Transcriptional Level.
Comparatively little is known about the transcription of type 1 fimbrial structural genes, but as described above, the recombinases inhibit PfimA. Perhaps not surprisingly, many of the factors that affect the fim inversion, including IHF, H-NS, and RpoS, also affect PfimA (14, 52, 171). Not only do mutants lacking IHF phase vary at much lower frequencies than wild-type E. coli cells, but the transcription of the fimbrial structural genes is decreased markedly as well (52). The mutation of IHF site 2 decreases PfimA activity, and hence, the effect is apparently direct (16). Several groups have reported an effect of H-NS on PfimA activity. However, whereas in an early study H-NS was reported to be a repressor later work indicated that it has the opposite effect (51, 171). Differences in strain background or in growth conditions may explain these differences (14).
Phase Variation in a fimS Locked-On Background.
A surprising finding is that mutants that contain fimS locked in the on orientation continue to show phase-variable expression of type 1 fimbriae (124). The basis for this has not been established, but it cannot be explained by variation in the activity of PfimA since the effect is seen even when fimS is replaced by the tac promoter. Thus, the effect apparently reflects control of type 1 fimbriation at a step subsequent to the initiation of transcription of the fimbrial structural genes.
Regulation of CS18 Fimbriae by Site-Specific Recombination.
While many fimbrial operons are regulated by methylation, site-specific recombination in E. coli has appeared to be restricted to a regulatory mechanism for type 1 fimbriation. However, the phase variation of CS18 fimbriae of ETEC was shown recently to be controlled by a comparable inversion-dependent mechanism (88). The expression of the plasmid-encoded fimbriae is, like that of fim, determined by the orientation of a short invertible element that lies immediately upstream of the fimbrial assembly and structural subunits. The fot invertible region comprises a 280-bp segment of DNA flanked by two 16-bp inverted repeats and, hence, is close in size to fimS. In addition, the inversion of the fot invertible element requires one of the two recombinases, FotS and FotT. The fot and fim recombinases show around 50% identity, including the highly conserved tetrad of amino acids essential for catalysis by lambda integrase and related tyrosine recombinases. Although not apparently controlled by either IHF or Lrp, the two fot recombinases seem to have roles parallel to those of FimB (FotS) and FimE (FotT). Furthermore, the 3' end of fotT overlaps the left inverted repeat of the invertible element, and fotT may, like fimE, be controlled differentially in on- and off-phase cells.
Sequence analysis in combination with studies on the regulation of the expression of the P pili encoded by the pap operon in E. coli led to the identification of a family of fimbrial operons that share basic principles underlying the molecular mechanism of phase variation (194). Except for the pef operon of Salmonella serovar Typhimurium, all members identified have been from E. coli isolates. The most extensively studied is pap, which provides the paradigm for members of this group and is discussed herein. Our treatment of pap is followed by a brief discussion of variations found among other members of this regulatory family. It should be emphasized that the operons of the pap regulatory family can be members of different families that are categorized based on the biochemical properties of the major fimbrial subunits.
The expression of the pap operon is controlled by numerous factors. A key element is the phase variation mechanism that determines if a cell can express Pap pili (the epigenetic on phase) or not (the epigenetic off phase) (120), which is regulated at the level of transcription (19). Epigenetic regulation in bacteria, including pap phase variation, was recently reviewed in detail (34). Here, the key elements of the phase variation mechanism and the way that the CpxAR regulatory system, a two-component regulatory system that senses envelope stress, affects this mechanism are outlined. Additional regulatory factors that are required for either transcriptional activation or repression but that do not directly affect the epigenetic switch are summarized next, and finally, posttranscriptional regulation is addressed. The conclusions and the model for the regulation of pap expression that are presented here are based on extensive analyses using a wide variety of experimental approaches. These include analyses of pap expression and phase variation in bacterial populations, including isolates with mutations in the pap regulatory region or in regulatory genes. These in vivo assays have been complemented by in vitro analyses of DNA binding sites and of DNA-protein binding constants and in vitro transcription assays with both wild-type and mutant DNA sequences.
Roles of Lrp, Dam, and the pap GATC Sequences.
The main promoter for the pap operon, the papBA promoter (PpapBA), lies in an intergenic region between the genes encoding the regulatory proteins PapI and PapB (Fig. 6) (7). Key regulatory elements in this intergenic region are two GATC target sequences for deoxyadenosine methyltransferase (Dam) and binding sites for the global regulatory protein Lrp. The GATC sequences are spaced 102 bp apart and are designated GATCdist, which has also been referred to as GATC-I or GATC1028, and GATCprox, which has also been referred to as GATC-II or GATC1130. In addition to binding sites for Lrp, there are binding sites for the regulatory proteins cyclic AMP (cAMP)-catabolite activator protein (CAP) (72) and CpxR (80), and H-NS binds to this intergenic region as well (209).
The regulatory region contains six binding sites for the global regulator Lrp (18, 19, 22, 23, 24). The binding of Lrp is cooperative, such that binding occurs simultaneously at sites 1, 2, and 3 or at sites 4, 5, and 6 (141, 142). Lrp binding is mutually exclusive, and thus, all six sites rarely appear to be occupied simultaneously in vivo (79, 81). The role of Lrp in regulating transcription from PpapBA depends on the sites where it is bound. Lrp bound at sites 1, 2, and 3 functions as a repressor of PpapBA transcription, whereas when it is bound at sites 4, 5, and 6, it functions as an activator (196, 204) (Fig. 2). However, Lrp requires PapI to bind at sites 4, 5, and 6 (see below).
GATCdist is contained within Lrp binding site 5, and GATCprox is within Lrp binding site 2 (23, 24). Lrp binding blocks the access of Dam to the GATC sequence contained in the occupied site, resulting in an unmethylated GATC sequence (22, 23, 24, 193, 204). The 3 bp flanking the GATC sequences may also contribute to a relatively low methylation rate (151), facilitating this Lrp-dependent methylation protection. This methylation protection occurs despite the fact that Dam is constitutively expressed in the cell. The result of this protection combined with mutually exclusive binding is that on and off cells have characteristic patterns of methylation of the GATC sequences in the pap regulatory region (Fig. 6) (18, 204).
The differential methylation pattern of the pap GATC sequences is not just an in vivo footprint of Lrp binding but is an essential feature for pap phase variation. This is because this methylation decreases the Lrp binding affinity for its pap sequences (23, 142). The methylation of GATCprox is essential for maintaining the protein binding pattern of the on phase by decreasing the affinity of the Lrp-PapI complex at sites 1, 2, and 3 (142). The methylation of GATCdist is required for stabilizing the nucleoprotein complex of the off phase by decreasing the affinity of Lrp as well as Lrp-PapI for sites 4, 5, and 6 (78, 79, 142). Thus, the differential methylation states play a role in reestablishing the protein binding patterns and thus contribute to the inheritance of the pap expression phase.
PapB, PapI, and the Autoregulatory Loop.
Phase variation and the transcription of PpapBA require the 8-kDa PapI protein (102, 141). The papI gene is located upstream of PpapBA and is transcribed from the divergent promoter PpapI. PapI binds to DNA by itself but with such low affinity that binding cannot be readily detected (81). More importantly, PapI is found in a complex with Lrp but only when Lrp is bound at pap. This specific interaction can be attributed to a PapI response element that consists of the sequence CGATC (81). This finding, in addition to the identification of TTT as significant for Lrp binding (141), explains the conservation of the sequences surrounding the two Dam target sites, also referred to as GATC boxes, among the pap family (192).
The presence of PapI results in an increased affinity of Lrp for site 5, and thus, PapI facilitates the switch from off to on (81, 141). Through the direct activation of PpapI, PapI levels increase, and this increase facilitates the maintenance of the on phase (79). Furthermore, the transcription of PpapI is activated by low concentrations of PapB, encoded by the first gene transcribed from PpapBA. This positive feedback between the two promoters is an essential element of the maintenance or self-perpetuation of the on phase (78, 79). However, this activation loop is controlled, since high concentrations of PapB repress transcription at the PpapBA promoter and thus lower the PapB expression level (7, 212, 214). In addition, high levels of PapI enhance the affinity of Lrp for site 2. The binding of Lrp at sites 1, 2, and 3 leads to PpapBA repression and, through mutual exclusion, a lack of activation of both PpapI and PpapBA.
Switch in Expression State.
The switch frequency for pap-17 expression in minimal medium with glycerol as the carbon and energy source is about 1 in 100 cells per generation for the on-to-off state and 1 in 10,000 cells per generation for the reverse direction (19, 120). To obtain a switch between the nucleoprotein complexes required for the on and off positions, an altered outcome of competition between Dam and Lrp for the two GATC-containing binding sites in pap is required (23, 79, 81, 192). For this alteration to occur, the methylation state of the GATC sequences must change. Thus, it is predicted that DNA replication is required for two reasons (79). First, the passage of the replication fork disrupts existing nucleoprotein complexes. Second, and equally important, the associated DNA synthesis generates DNA that is temporarily hemimethylated, which in the absence of a demethylating enzyme in E. coli provides a significant mechanism for obtaining hemi- and nonmethylated GATC sequences.
Detailed analyses of the binding affinities of Lrp and Lrp-PapI for nonmethylated, hemimethylated, and methylated wild-type and mutated binding regions has led to the following model for the phase variation of pap (81). The on phase requires Lrp-PapI binding at sites 4, 5, and 6. Thus, for the switch from the off to the on phase, the methylated state of GATCdist has to change since it in essence blocks the binding of the Lrp-PapI complex by conferring decreased affinity of Lrp-PapI for sites 4, 5, and 6 (81, 141, 142, 204). However, the affinity of Lrp-PapI for sites 4, 5, and 6 with a hemimethylated GATCdist is higher (81), and thus, after DNA replication, an Lrp-PapI complex can bind to the GATCdist-containing region (7, 101, 196), initiating the switch. Interestingly, the affinity differs depending on whether the top or bottom strand contains the methyl group. This difference may cause the two daughter cells to have different chances of switching expression phase (81). Regardless, as a result of the binding of Lrp-PapI, the access of Dam to GATCdist is blocked, and after an additional round of DNA replication, a fully unmethylated GATCdist is obtained (204). Mutually exclusive binding will presumably contribute to the absence of Lrp at sites 1, 2, and 3 that allows the methylation of GATCprox. This, in turn, blocks the binding of the Lrp-PapI complex to the DNA region encompassing sites 1, 2, and 3, maintaining the on phase. The PapI- and PapB-dependent positive feedback loop then also contributes to the maintenance of the on phase.
Conversely, a switch from the on phase to the off phase is facilitated by a change from the methylated state of GATCprox after DNA replication to the hemimethylated state, allowing the binding of Lrp-PapI at sites 1, 2, and 3. The state is maintained as a result of new methylation at GATCdist (81). Interestingly, Lrp binds to sites 1, 2, and 3 regardless of the methylation state of GATCprox (142), supporting the observation that the off phase is the default phase in the absence of PapI.
CpxAR-Dependent Repression of the Switch.
Hung et al. identified the two-component CpxAR system as a regulator of pap expression (94). CpxAR is thought to signal envelope stress (45, 169, 180), but the precise signal is not known. Hung et al. showed that the activation of the CpxAR system results in an increase in pap transcription and the expression of Pap fimbriae (94). Hernday et al. also examined the role of CpxAR in pap expression but used a single-copy reporter system for pap expression instead of the multicopy system used by Hung et al. (80). Their results led to an opposite conclusion, namely, that the activation of CpxAR results in the repression of the transcription of pap and of the synthesis of Pap fimbriae. The discrepancy appears to derive from the use of the multi- or single-copy reporter system. The phase variation of pap is not apparent when the operon is present in multiple copies, and in pathogenic species, the pap operon exists as a single copy. At most, a single isolate contains only a few closely related operons (5, 75, 103, 139). Furthermore, the role of CpxAR as a repressor is dependent on the normal level of PapI (80), which is not maintained in a multicopy system. Taken together, these factors strongly indicate that the findings by Hernday et al. reflect the role of CpxAR in pap regulation in clinical isolates, and thus, their study is summarized further.
The activation of the CpxAR system results in the phosphorylation of CpxR (CpxR-P) (45). In vitro analyses showed that CpxR-P binds to sites both in and between the Lrp binding sites in a region spanning about 180 bp between PpapI and PpapBA. In vivo analyses of the pap GATC methylation states indicate that, in contrast to the binding of Lrp to the GATC regions, the binding of CpxR-P does not protect the sites from methylation. Also in contrast to Lrp, the methylation of the pap GATC sites does not affect the binding affinity of CpxR-P for the pap regulatory region. Analyses of in vivo methylation patterns indicate that CpxR-P does not displace Lrp bound at sites 1, 2, and 3 and thus does not affect pap regulation in cells in the off phase.
The role of CpxAR binding was examined both in vitro and in vivo, and the findings indicate that CpxAR activation blocks the off-to-on switch and, in addition, represses transcription from PpapBA (80). CpxR-P reduces the affinity of Lrp-PapI for sites 4, and 5, and 6 (80), and thus, CpxR-P may be an effective competitor for Lrp-PapI for binding at this region and thereby decrease the off-to-on frequency directly. This effect may be especially important with a hemimethylated GATCdist since the affinity of CpxR-P is not affected by the methylation (78, 80). In addition, the binding of CpxR-P at the vacant region encompassing sites 1, 2, and 3 in on-phase cells may repress the transcription of PpapBA. This repression would result directly in decreased expression and, through the PapB-PapI feedback loop, also lead to an increased frequency of switching to the off phase. Interestingly, CpxR-P is the first regulator to be identified that directly affects the pap switch frequency in response to environmental cues. This CpxAR-dependent regulation may provide feedback between P pilus assembly and the expression of the operon, since nonassembled units of the structural subunit PapE can activate the CpxAR system (115).
Additional Regulators of pap Transcription.
Other factors have been identified that affect the transcription of pap but do not directly affect the Lrp-, PapI-, and Dam-dependent switch, and these factors are therefore epistatic for the switch. An essential coactivator for the transcription of PpapBA is cAMP-catabolite regulator protein (CRP, or CAP), which mediates regulation in response to the carbon source. CRP binds between the GATCdist region and the PpapI promoter (7, 19, 61, 72, 203) (Fig. 6). Initial analyses by Forsman et al. using a multicopy pap coding system suggested that cAMP-CRP was an antirepressor of H-NS (61). However, analyses of phase variation and transcription using single-copy reporters and analyses of the effects of CRP activation mutations indicate that CRP functions as a direct activator of PpapBA. This function was confirmed with in vitro analyses of promoter activation (203), and the activation occurs despite the binding of CRP at the larger than usual distance of 215.5 bp from the activated promoter (203). cAMP-CRP may play a dual role in pap expression since the complex bound at this same site also activates expression from PpapI (72). However, whether this is a direct effect on PpapI or an indirect effect on PpapBA that is mediated through PapB remains to be determined. This dependency on CRP couples Pap expression to the available carbon source and the nutritional state of the bacterium.
The effect of other environmental conditions on pap expression was examined in a comparative study (209). The repression of pap expression at low temperatures (23°C or below), in medium with glucose, in rich medium, and at high osmolarity was observed, and repression mediated by these factors was completely or partially relieved by an hns mutation (209). H-NS is not essential for pap phase variation but can repress transcription from PpapBA in the absence of Lrp (196), and at 37°C, H-NS modulates the pap switch frequency, as evidenced by a two- to threefold reduction in the off-to-on rate in an hns mutant (205). H-NS can bind to the pap intergenic region and protect both GATC sequences from methylation. This binding also appears to block transcription directly, since after a temperature downshift, transcription ceases before the off DNA methylation pattern is observed (205). Mutational analyses also identified rimJ as a gene that contributes to the thermoregulation of pap (207, 208), and like the hns mutation, a rimJ mutation relieves the repression of pap expression mediated by low temperatures. The rimJ mutation results in an increase of cells in the on phase. RimJ is annotated as an N-terminal acetylase of the ribosomal protein S5, and how it controls pap expression remains to be elucidated (206). Mutations in both hns and rimJ also relieve repression by exogenous glucose and rich medium.
Posttranscriptional Regulation of pap.
The activation of PpapBA results in the synthesis of a polycistronic transcript. It appears that a significant percentage of the transcripts terminate after papA so that the level of transcripts with papH to papG is significantly lower than the level of transcripts with papA, corresponding to a lower level of the minor subunits than of PapA (7). It is not certain, however, that there are no additional, minor promoters downstream of papA. The major transcript includes papA and papB (7), yet PapA is also expressed at a much higher level than PapB. This can, in part at least, be attributed to RNase E-dependent posttranscriptional processing at a sequence in the intergenic region between the papA and papB cistron (6, 138). The cleavage step is the rate-limiting step defining the stability, and the difference in transcript stability is responsible for the difference in the levels of expression of PapA and PapB (138). The removal of the mRNA processing sequences results in decreased PapA expression and the synthesis of short fimbriae (137).
Regulation of Expression among Phase-Variable Members of the pap Family.
Members of the pap-like (regulatory) family of fimbrial operons are identified based on sequence conservation of the regulatory elements and regulatory proteins only. Specifically, conservation of the GATC regions (GATC boxes), as described above, and the Lrp binding sites exists, and each operon encodes homologs of the PapI and PapB proteins.
In E. coli, the members of this family, including the prf, daa, and sfa operons, all seem to have retained the basic pap-like organization of the regulatory region. It has been shown previously that the expression of sfa (194, 197), daa (12, 194, 197), clp (42) (122), and foo (10, 44, 77, 121), encoding, respectively, the Sfa, F1845, CS31A, and F165(1) adhesins, phase varies in a Dam- and Lrp-dependent manner similar to that of pap. F165(2), very closely related to F165(1), also phase varies (44). Unfortunately, the operon coding for F165(2) was designated fot by Daigle et al. (44) even though fot was used previously to designates the CS18-encoding operon, discussed in "Operons regulated by site-specific recombination," above (88, 200). Therefore, we suggest that the operon coding for F165(2) be renamed foo-2. Compared to the elements and genes in all these pap-like operons, those in the Salmonella pef operon have been rearranged (see below).
Despite the general conservation of operon structure among the E. coli operons of the pap family, the roles of the homologs of PapI vary. Whereas the cognate regulator is a positive regulator of expression for daa, sfa, and foo, it is a negative regulator for pef (136) and clp (122). Minor sequence differences that result in variations in switch frequencies and levels of expression even between closely related operons occur. For example, pap-17 has a 15-fold-higher on-to-off switch frequency than pap-21 (19), which possibly can be attributed to an additional PapB binding site in the regulatory region (212). Sequence variations affecting the rate of methylation by Dam (194) may also contribute to differences in switch frequencies (151).
Differential responses to environmental conditions among the members of the pap family also occur. For example, this is evident in the effect of leucine on the expression of some of these operons. Leucine is known to affect Lrp binding to its cognate sites, but the magnitude of the effect depends both on the Lrp concentration and on the affinity for the site (20, 31, 59). The latter can be affected by minor sequence variations in the binding sites. Pap expression, for example, is not affected by leucine (24), but leucine represses the expression of the CS31A-encoding operon clp (122) and of the F165(1)-encoding operon foo (44, 77). In previous studies, leucine appeared to affect both the phase variation and the transcription of foo (10, 77), which are mediated by Lrp, even though a later report indicated that phase variation is not affected (43). Crost et al. found that leucine does not appear to change the DNA methylation pattern at the GATC sequences of clp, suggesting that Lrp may not be involved in this regulation. In contrast, alanine does affect the methylation state (42). Like that of pap, the expression of foo and clp decreases as a result of exogenous glucose and low temperatures (42, 43, 122).
The expression of daa responds in general to the same factors as that of pap. An hns mutation relieves repression by low temperatures similar to that of pap and completely relieves repression by high osmolarity and rich medium (206, 209). In contrast to one in pap, however, a mutation in hns results in an 11.5-fold increase in the off-to-on switch rate under nonrepressing conditions (209). Also, mutations in rimJ do not affect the environmental regulation of daa expression (206).
The only member of this family in Salmonella is the pef operon. This operon contains regulatory sequences and proteins that place it in the pap family of fimbrial operons, and the principles underlying the phase variation regulatory mechanism are shared with those for pap. Nevertheless, pef regulation has distinctive properties, including the addition of a third Dam target sequence in the regulatory region and the positioning of the gene coding for the PapI homolog PefI downstream of the structural genes (136). PefI results in the repression of transcription and contributes to the Lrp-dependent methylation protection of a different GATC site than that of pap. Previous analyses have revealed distinct differences in the correlation between DNA methylation states and fimbrial expression, which is less stringent for pef than for pap regulation. Furthermore, pef expression is induced at low pHs, which has not been reported for the other pap family members. Similar to the E. coli operons, pef is repressed at low temperatures and by H-NS, but it is also repressed by the stationary-phase sigma factor RpoS (136).
Posttranscriptional Regulation in the pap Family.
The operons with gene organization patterns similar to that of pap must also have mechanisms to ensure the correct relative amounts of production of the various subunits. This issue has been investigated in detail for sfa and daa transcripts. The overall organization of the sfa cluster is very similar to that of pap (172), with a strong promoter before sfaB. The main difference is that there are only three, not six, genes in the distal part of the operon. Analogous to pap transcripts, the transcript for sfaA is processed by RNase E from a precursor sfaBA transcript, and sequences similar to that in pap are found at the processing site. This, probably in combination with partial transcription termination at the 3' end of sfaA and differential mRNA stability, results in an abundance of sfaA transcripts (128, 137). However, additional endoribonucleolytic sites in the full-length transcript exist, and a total of seven transcripts, including those of sfaGSH, representing three minor subunits, have been identified (8). A biochemical analysis of pili indicates that these three proteins are indeed present in equimolar amounts (8).
In the daa operon encoding F1485 pili, the organization of the genes is quite different. The adhesin- and major subunit-encoding daaE is not adjacent to the papB homolog daaA but is the fifth and final gene of the daaABCDPE transcript (12). Processing of the transcript and differential stabilities of the cleavage products also result in a relative accumulation of the daaE transcript (11, 13). However, none of the major endonucleases, RNase E, III, and P, is involved (11, 118). The required processing site is located in daaP, encoding a small, 57-amino-acid peptide (118, 119). This processing is directly coupled to the translation of daaP (118), and based on a mutational analysis of the coding sequence and the ribosome binding site of daaP, it was shown that in cis, the translation of daaP is required for processing to occur. It appears that the translation of the entire gene is required, but as yet only the absence of translation of a specific tripeptide sequence (Gly-Pro-Pro) at the C terminus of DaaP can be shown to decrease processing (118, 119). Mutants defective in processing were isolated previously, and one mutation was mapped to and may be complemented by hrpA, indicating that the gene product, a putative DEAH-box RNA helicase, is involved in the processing of the daa transcript. However, an hrpA mutation did not completely abolish processing, so additional factors may be required (109). Future work will have to identify the precise mechanisms and roles of daaP translation and HrpA in daa transcript processing.
pap
Family Operons That Are Not Phase Variable.
F4 (K88) and F5 (K99) fimbriae are important virulence factors in porcine ETEC (134). The F4-encoding operon fae is included in the pap regulatory family based on the presence of genes for the PapI and PapB homologs FaeA and FaeB. The regulatory region also contains GATC sequences, Lrp and Dam contribute to fae regulation, Lrp-dependent protection of the GATC sequences in the regulatory region occurs, and FaeA appears to act like PapI in modulating Lrp binding (89, 90). The PapB homolog FaeB does not appear to contribute to regulation. However, fae expression is not controlled by phase variation, and FaeA is a negative regulator of fae expression (89, 90). These differences can be attributed at least in part to insertion sequence elements in the regulatory region that disrupt the putative feedback loop between the faeA and faeB promoters and to other sequence variations that include a third GATC sequence (references 89 and 90; reviewed in reference 198). The regulation of fae is discussed in more detail in Chapter (Adhesins of Enterotoxigenic Escherichia coli Strains That Infect Animals).
The fan operon encodes F5 (K99) fimbriae in porcine ETEC (134). The fan regulatory region has limited homology to pap, and fan regulation has relatively little in common with pap regulation. This operon encodes two PapB homologs, FanA and FanB, but no PapI homolog, and Dam is not involved in regulating expression. The expression of fan is Lrp dependent and leucine sensitive but does not phase vary (24, 95, 167, 168). This illustrates that regulatory modules can diverge over time, leading to variations on a regulatory principle, and that care must be taken in predicting regulatory effects and mechanisms based on the presence of pap-like regulatory sequences and regulatory proteins.
The previous sections described the control of fimbrial gene expression by coordinate actions of global regulatory factors and those encoded by the fimbrial operons themselves. The global regulatory factors include proteins, small RNAs, and nucleotides. Many of these regulate more than one of the fimbrial operons, thus establishing the coordinated expression of a subset of fimbriae in response to specific environmental stimuli. The operon-encoded, or local, regulators were long considered to act only upon the cognate operon. However, many years ago Nowicki et al. found that strains very rarely present more than one type of fimbria on the surface at a given time (143), and recent studies have determined that this property can be brought about by some local regulators’ mediating the cross regulation of other fimbrial operons. Rather than being merely accidental, this extended regulation has likely evolved to be of benefit to the bacterium and a key factor to consider when investigating fimbrial regulation, and therefore, it is referred to herein as cross regulation.
Within the pap regulatory family, the PapI- and PapB-like proteins show a high degree of homology, and this homology allows some cross regulation to occur among members of this family. For example, functional complementation of sfaC mutations is obtained with papI (71) and that of papI mutations is obtained with daaF (197), and expected Dam-Lrp-DNA interactions mediated by PapI and daa regulatory DNA have occurred in vitro (197). Thus, the feedback loops mediated by the PapB- and PapI-like regulatory proteins can control the expression of noncognate members of the pap regulatory family. UPEC isolates and especially pyelonephritis-associated isolates contain more than one Pap-related gene cluster (86, 120, 127, 202), and therefore, this cross regulation can be relevant for host-pathogen interactions. Many E. coli genomes encode multiple fimbrial operons of the pap family, but the exact contribution of individual fimbrial operons to virulence and the relevance of any cross regulation remains to be determined (41, 74, 117, 178, 179, 210, 211).
PapB can also lead to coordinated expression that extends beyond the pap family, including the type 1 fimbrial expression in E. coli (84, 86, 87, 213). Specifically, it was shown previously that the expression of PapB from a multicopy plasmid affects the expression of the fim operon encoding type1 fimbriae. This cross regulation may involve two pathways. First, the expression of the FimE recombinase is directly affected by the expression of PapB, resulting in an increased on-to-off switch frequency. Second, FimB-mediated recombination is inhibited by PapB in vitro. The mechanism needs to be elucidated, but PapB binding occurs at the fimS region (87, 213), and two PapB amino acids, Leu-82 and Ile-81 in the C terminus of PapB, are essential (87). This pap-fim cross regulation is likely to be relevant in vivo since it occurs in many UPEC clinical isolates, as determined by analyses of type 1 expression at the single-cell and population levels and of the orientation of the fimS element (86). Consistent with the requirement of the two PapB amino acids, cross regulation can also be exerted by homologs SfaB and the Salmonella PefB protein (87) but not by the homologs DaaA (84), FanA, FanB, FaeB, and ClpB (87).
Snyder et al. showed that the converse of the regulation described above is also true: fim expression results in the downregulation of pap expression (178). This downregulation was detected by using microarrays to examine gene expression in the uropathogenic isolate E. coli CFT073, but how this regulation is effected is not known. Snyder et al. also found that F1C fimbriae, which are of the pap regulatory family, that are not expressed by the wild-type isolate are expressed when Pap and type 1 fimbrial production is abolished by the deletion of the respective structural fimbrial genes. In this mutant, the pap regulatory and fim recombinase genes are still present, and therefore, this cross regulation may not involve fimbrial regulatory proteins.
In Salmonella serovar Typhimurium, cross regulation between fimbrial operons and flagella occurs and, as Clegg and Hughes phrased it, leads to a decision to "stick or swim" (37). This cross regulation is mediated by the Salmonella serovar Typhimurium fim-encoded DNA binding protein FimZ, but again the mechanism is not clear (37). In Proteus mirabilis, the expression of mrpJ of the mannose-resistant Proteus-like fimbrial operon can repress the transcription of the flagellar operon flhDC, probably indirectly. An mrpJ mutation in P. mirabilis can be complemented by the expression of papX from the E. coli pap operon (116), suggesting that coordinated expression of flagella and fimbriae, specifically of the pap family, may also occur in E. coli. Finally, cross regulation may extend even to metabolism, as indicated by the role of CsgD, the regulatory protein of the curli operon, in the expression of cellulose production (70, 164, 165, 166), and therefore the control of biofilm formation by many strains of Salmonella serovars Typhimurium and Enteritidis (46).
These examples illustrate that the regulators encoded by the fimbrial operons can have a significant effect on the bacterial phenotype and behavior by controlling the expression of genes outside of the cognate operon. Other examples of possible regulatory networks and complex bacterial behavior are reviewed in reference 85. As more of these networks are being scrutinized, it seems likely that additional cross regulation between fimbrial operons and other cellular genes will be identified. The biological relevance of the complex regulatory networks points to a benefit of coordinating the temporal and spatial production of fimbriae during infection, possibly to control tissue tropism, immune evasion, and dispersal.
This chapter illustrates how fimbrial expression is subject to elaborate control by both local and global regulatory circuits. Novel principles undoubtedly remain to be elucidated, however, and as this process progresses, this chapter will continue to be a valuable springboard for the discussion of this topic. The realization that fimbriae elicit innate host immune responses adds particular relevance to these endeavors, offering to enhance considerably our understanding of the host-parasite relationship in both commensal and pathogenic interactions.
We thank M. Goulian and D. Schifferli for helpful discussions.
This work was supported by grant 076360/Z/05/Z from the Wellcome Trust to I.B.; work in the M.v.d.W. lab is supported by grant BB/C502849/1 from the BBSRC and grant MIRG-7-CT-2005-017104 from the European Union.
References
1. Aberg, A., V. Shingler, and C. Balsalobre. 2006. (p)ppGpp regulates type 1 fimbriation of Escherichia coli by modulating the expression of the site-specific recombinase FimB. Mol. Microbiol. 60:1520–1533.[PubMed] [CrossRef]
2. Abraham, J. M., C. S. Freitag, J. R. Clements, and B. I. Eisenstein. 1985. An invertible element of DNA controls phase variation of type 1 fimbriae of Escherichia coli. Proc. Natl. Acad. Sci. USA 82:5724–5727.[PubMed] [CrossRef]
3. Arnqvist, A., A. Olsen, and S. Normark. 1994. Sigma S-dependent growth-phase induction of the csgBA promoter in Escherichia coli can be achieved in vivo by sigma 70 in the absence of the nucleoid-associated protein H-NS. Mol. Microbiol. 13:1021–1032.[PubMed] [CrossRef]
4. Arnqvist, A., A. Olsen, J. Pfeifer, D. G. Russell, and S. Normark. 1992. The Crl protein activates cryptic genes for curli formation and fibronectin binding in Escherichia coli HB101. Mol. Microbiol. 6:2443–2452.[PubMed]
5. Arthur, M., C. Campanelli, R. D. Arbeit, C. Kim, S. Steinbach, C. E. Johnson, R. H. Rubin, and R. Goldstein. 1989. Structure and copy number of gene clusters related to pap P-adhesin operon of uropathogenic Escherichia coli. Infect. Immun. 57:314–321.[PubMed]
6. Baga, M., M. Goransson, S. Normark, and B. E. Uhlin. 1988. Processed mRNA with differential stability in the regulation of E. coli pilin gene expression. Cell 52:197–206.[PubMed] [CrossRef]
7. Baga, M., M. Goransson, S. Normark, and B. E. Uhlin. 1985. Transcriptional activation of a pap pilus virulence operon from uropathogenic Escherichia coli. EMBO J. 4:3887–3893.[PubMed]
8. Balsalobre, C., J. Morschhauser, J. Jass, J. Hacker, and B. E. Uhlin. 2003. Transcriptional analysis of the sfa determinant revealing multiple mRNA processing events in the biogenesis of S fimbriae in pathogenic Escherichia coli. J. Bacteriol. 185:620–629.[PubMed] [CrossRef]
9. Barnhart, M. M., J. Lynem, and M. R. Chapman. 2006. GlcNAc-6P levels modulate the expression of curli fibers by Escherichia coli. J. Bacteriol. 188:5212–5219.[PubMed] [CrossRef]
10. Berthiaume, F., C. Crost, V. Labrie, C. Martin, E. B. Newman, and J. Harel. 2004. Influence of L-leucine and L-alanine on Lrp regulation of foo, coding for F1651, a Pap homologue. J. Bacteriol. 186:8537–8541.[PubMed] [CrossRef]
11. Bilge, S. S., J. M. Apostol, Jr., M. A. Aldape, and S. L. Moseley. 1993. mRNA processing independent of RNase III and RNase E in the expression of the F1845 fimbrial adhesin of Escherichia coli. Proc. Natl. Acad. Sci. USA 90:1455–1459.[PubMed] [CrossRef]
12. Bilge, S. S., J. M. Apostol, Jr., K. J. Fullner, and S. L. Moseley. 1993. Transcriptional organization of the F1845 fimbrial adhesin determinant of Escherichia coli. Mol. Microbiol. 7:993–1006.[PubMed] [CrossRef]
13. Bilge, S. S., C. R. Clausen, W. Lau, and S. L. Moseley. 1989. Molecular characterization of a fimbrial adhesin, F1845, mediating diffuse adherence of diarrhea-associated Escherichia coli to HEp-2 cells. J. Bacteriol. 171:4281–4289.[PubMed]
14. Blomfield, I. C. 2001. The regulation of pap and type 1 fimbriation in Escherichia coli. Adv. Microb. Physiol. 45:1–49.[PubMed] [CrossRef]
15. Blomfield, I. C., P. J. Calie, K. J. Eberhardt, M. S. McClain, and B. I. Eisenstein. 1993. Lrp stimulates phase variation of type 1 fimbriation in Escherichia coli K-12. J. Bacteriol. 175:27–36.[PubMed]
16. Blomfield, I. C., D. H. Kulasekara, and B. I. Eisenstein. 1997. Integration host factor stimulates both FimB- and FimE-mediated site-specific DNA inversion that controls phase variation of type 1 fimbriae expression in Escherichia coli. Mol. Microbiol. 23:705–717.[PubMed] [CrossRef]
17. Blomfield, I. C., M. S. McClain, J. A. Princ, P. J. Calie, and B. I. Eisenstein. 1991. Type 1 fimbriation and fimE mutants of Escherichia coli K-12. J. Bacteriol. 173:5298–5307.[PubMed]
18. Blyn, L. B., B. A. Braaten, and D. A. Low. 1990. Regulation of pap pilin phase variation by a mechanism involving differential dam methylation states. EMBO J. 9:4045–4054.[PubMed]
19. Blyn, L. B., B. A. Braaten, C. A. White-Ziegler, D. H. Rolfson, and D. A. Low. 1989. Phase-variation of pyelonephritis-associated pili in Escherichia coli: evidence for transcriptional regulation. EMBO J. 8:613–620.[PubMed]
20. Borst, D. W., R. M. Blumenthal, and R. G. Matthews. 1996. Use of an in vivo titration method to study a global regulator: effect of varying Lrp levels on expression of gltBDF in Escherichia coli. J. Bacteriol. 178:6904–6912.[PubMed]
21. Bougdour, A., C. Lelong, and J. Geiselmann. 2004. Crl, a low temperature-induced protein in Escherichia coli that binds directly to the stationary phase sigma subunit of RNA polymerase. J. Biol. Chem. 279:19540–19550.[PubMed] [CrossRef]
22. Braaten, B. A., L. B. Blyn, B. S. Skinner, and D. A. Low. 1991. Evidence for a methylation-blocking factor (mbf) locus involved in pap pilus expression and phase variation in Escherichia coli. J. Bacteriol. 173:1789–1800.
23. Braaten, B. A., X. Nou, L. S. Kaltenbach, and D. A. Low. 1994. Methylation patterns in pap regulatory DNA control pyelonephritis-associated pili phase variation in E. coli. Cell 76:577–588.[PubMed] [CrossRef]
24. Braaten, B. A., J. V. Platko, M. W. van der Woude, B. H. Simons, F. K. de Graaf, J. M. Calvo, and D. A. Low. 1992. Leucine-responsive regulatory protein controls the expression of both the pap and fan pili operons in Escherichia coli. Proc. Natl. Acad. Sci. USA 89:4250–4254.[PubMed] [CrossRef]
25. Brinton, C. C., Jr. 1959. Non-flagellar appendages of bacteria. Nature 183:782–786.[PubMed] [CrossRef]
26. Brombacher, E., A. Baratto, C. Dorel, and P. Landini. 2006. Gene expression regulation by the curli activator CsgD protein: modulation of cellulose biosynthesis and control of negative determinants for microbial adhesion. J. Bacteriol. 188:2027–2037.[PubMed] [CrossRef]
27. Brombacher, E., C. Dorel, A. J. Zehnder, and P. Landini. 2003. The curli biosynthesis regulator CsgD co-ordinates the expression of both positive and negative determinants for biofilm formation in Escherichia coli. Microbiology 149:2847–2857.[PubMed] [CrossRef]
28. Brown, P. K., C. M. Dozois, C. A. Nickerson, A. Zuppardo, J. Terlonge, and R. Curtiss III. 2001. MlrA, a novel regulator of curli (AgF) and extracellular matrix synthesis by Escherichia coli and Salmonella enterica serovar Typhimurium. Mol. Microbiol. 41:349–363.[PubMed] [CrossRef]
29. Bryan, A., P. Roesch, L. Davis, R. Moritz, S. Pellett, and R. A. Welch. 2006. Regulation of type 1 fimbriae by unlinked FimB- and FimE-like recombinases in uropathogenic Escherichia coli strain CFT073. Infect. Immun. 74:1072–1083.[PubMed] [CrossRef]
30. Burns, L. S., S. G. Smith, and C. J. Dorman. 2000. Interaction of the FimB integrase with the fimS invertible DNA element in Escherichia coli in vivo and in vitro. J. Bacteriol. 182:2953–2959.[PubMed] [CrossRef]
31. Calvo, J. M., and R. G. Matthews. 1994. The leucine-responsive regulatory protein, a global regulator of metabolism in Escherichia coli. Microbiol. Rev. 58:466–490.[PubMed]
32. Caron, J., L. M. Coffield, and J. R. Scott. 1989. A plasmid-encoded regulatory gene, rns, required for expression of the CS1 and CS2 adhesins of enterotoxigenic Escherichia coli. Proc. Natl. Acad. Sci. USA 86:963–967.[PubMed] [CrossRef]
33. Caron, J., and J. R. Scott. 1990. An rns-like regulatory gene for colonization factor antigen I (CFA/I) that controls expression of CFA/I pilin. Infect. Immun. 58:874–878.[PubMed]
34. Casadesus, J., and D. Low. 2006. Epigenetic gene regulation in the bacterial world. Microbiol. Mol. Biol. Rev. 70:830–856.[PubMed] [CrossRef]
35. Chirwa, N. T., and M. B. Herrington. 2003. CsgD, a regulator of curli and cellulose synthesis, also regulates serine hydroxymethyltransferase synthesis in Escherichia coli K-12. Microbiology 149:525–535.[PubMed] [CrossRef]
36. Clegg, S., L. S. Hancox, and K. S. Yeh. 1996. Salmonella typhimurium fimbrial phase variation and FimA expression. J. Bacteriol. 178:542–545.[PubMed]
37. Clegg, S., and K. T. Hughes. 2002. FimZ is a molecular link between sticking and swimming in Salmonella enterica serovar Typhimurium. J. Bacteriol. 184:1209–1213.[PubMed] [CrossRef]
38. Clouthier, S. C., S. K. Collinson, A. P. White, P. A. Banser, and W. W. Kay. 1998. tRNAArg (fimU) and expression of SEF14 and SEF21 in Salmonella enteritidis. J. Bacteriol. 180:840–845.[PubMed]
39. Collinson, S. K., S. C. Clouthier, J. L. Doran, P. A. Banser, and W. W. Kay. 1996. Salmonella enteritidis agfBAC operon encoding thin, aggregative fimbriae. J. Bacteriol. 178:662–667.[PubMed]
40. Condemine, G., C. Berrier, J. Plumbridge, and A. Ghazi. 2005. Function and expression of an N-acetylneuraminic acid-inducible outer membrane channel in Escherichia coli. J. Bacteriol. 187:1959–1965.[PubMed] [CrossRef]
41. Connell, I., W. Agace, P. Klemm, M. Schembri, S. Marild, and C. Svanborg. 1996. Type 1 fimbrial expression enhances Escherichia coli virulence for the urinary tract. Proc. Natl. Acad. Sci. USA 93:9827–9832.[PubMed] [CrossRef]
42. Crost, C., A. Garrivier, J. Harel, and C. Martin. 2003. Leucine-responsive regulatory protein-mediated repression of clp (encoding CS31A) expression by L-leucine and L-alanine in Escherichia coli. J. Bacteriol. 185:1886–1894.[PubMed] [CrossRef]
43. Crost, C., J. Harel, F. Berthiaume, A. Garrivier, M. C. Tessier, H. Rakotoarivonina, and C. Martin. 2004. Influence of environmental cues on transcriptional regulation of foo and clp coding for F165(1) and CS31A adhesins in Escherichia coli. Res. Microbiol. 155:475–482.[PubMed] [CrossRef]
44. Daigle, F., C. Forget, C. Martin, M. Drolet, M. C. Tessier, H. Dezfulian, and J. Harel. 2000. Effects of global regulatory proteins and environmental conditions on fimbrial gene expression of F165(1) and F165(2) produced by Escherichia coli causing septicaemia in pigs. Res. Microbiol. 151:563–574.[PubMed] [CrossRef]
45. Danese, P. N., W. B. Snyder, C. L. Cosma, L. J. Davis, and T. J. Silhavy. 1995. The Cpx two-component signal transduction pathway of Escherichia coli regulates transcription of the gene specifying the stress-inducible periplasmic protease, DegP. Genes Dev. 9:387–398.[PubMed] [CrossRef]
46. Da Re, S., and J. M. Ghigo. 2006. A CsgD-independent pathway for cellulose production and biofilm formation in Escherichia coli. J. Bacteriol. 188:3073–3087.[PubMed] [CrossRef]
47. de Haan, L. A., G. A. Willshaw, B. A. van der Zeijst, and W. Gaastra. 1991. The nucleotide sequence of a regulatory gene present on a plasmid in an enterotoxigenic Escherichia coli strain of serotype O167:H5. FEMS Microbiol. Lett. 67:341–346.[PubMed] [CrossRef]
48. Donato, G. M., and T. H. Kawula. 1999. Phenotypic analysis of random hns mutations differentiate DNA-binding activity from properties of fimA promoter inversion modulation and bacterial motility. J. Bacteriol. 181:941–948.[PubMed]
49. Donato, G. M., M. J. Lelivelt, and T. H. Kawula. 1997. Promoter–specific repression of fimB expression by the Escherichia coli nucleoid–associated protein H–NS. J. Bacteriol. 179:6618–6625.[PubMed]
50. Dorel, C., O. Vidal, C. Prigent-Combaret, I. Vallet, and P. Lejeune. 1999. Involvement of the Cpx signal transduction pathway of E. coli in biofilm formation. FEMS Microbiol. Lett. 178:169–175.[PubMed] [CrossRef]
51. Dorman, C. J., and N. N. Bhriain. 1992. Thermal regulation of fimA, the Escherichia coli gene coding for the type 1 fimbrial subunit protein. FEMS Microbiol. Lett. 78:125–130.[PubMed] [CrossRef]
52. Dorman, C. J., and C. F. Higgins. 1987. Fimbrial phase variation in Escherichia coli: dependence on integration host factor and homologies with other site-specific recombinases. J. Bacteriol. 169:3840–3843.[PubMed]
53. Dove, S. L., and C. J. Dorman. 1996. Multicopy fimB gene expression in Escherichia coli: binding to inverted repeats in vivo, effect on fimA gene transcription and DNA inversion. Mol. Microbiol. 21:1161–1173.[PubMed] [CrossRef]
54. Dove, S. L., and C. J. Dorman. 1994. The site-specific recombination system regulating expression of the type 1 fimbrial subunit gene of Escherichia coli is sensitive to changes in DNA supercoiling. Mol. Microbiol. 14:975–988.[PubMed] [CrossRef]
55. Edwards, R. A., and D. M. Schifferli. 1997. Differential regulation of fasA and fasH expression of Escherichia coli 987P fimbriae by environmental cues. Mol. Microbiol. 25:797–809.[PubMed] [CrossRef]
56. Eisenstein, B. I. 1981. Phase variation of type 1 fimbriae in Escherichia coli is under transcriptional control. Science 214:337–339.[PubMed] [CrossRef]
57. Eisenstein, B. I., D. S. Sweet, V. Vaughn, and D. I. Friedman. 1987. Integration host factor is required for the DNA inversion that controls phase variation in Escherichia coli. Proc. Natl. Acad. Sci. USA 84:6506–6510.[PubMed] [CrossRef]
58. El-Labany, S., B. K. Sohanpal, M. Lahooti, R. Akerman, and I. C. Blomfield. 2003. Distant cis-active sequences and sialic acid control the expression of fimB in Escherichia coli K-12. Mol. Microbiol. 49:1109–1118.[PubMed] [CrossRef]
59. Ernsting, B. R., J. W. Denninger, R. M. Blumenthal, and R. G. Matthews. 1993. Regulation of the gltBDF operon of Escherichia coli: how is a leucine-insensitive operon regulated by the leucine-responsive regulatory protein? J. Bacteriol. 175:7160–7169.[PubMed]
60. Ferrieres, L., and D. J. Clarke. 2003. The RcsC sensor kinase is required for normal biofilm formation in Escherichia coli K-12 and controls the expression of a regulon in response to growth on a solid surface. Mol. Microbiol. 50:1665–1682.[PubMed] [CrossRef]
61. Forsman, K., B. Sonden, M. Goransson, and B. E. Uhlin. 1992. Antirepression function in Escherichia coli for the cAMP-cAMP receptor protein transcriptional activator. Proc. Natl. Acad. Sci. USA 89:9880–9884.[PubMed] [CrossRef]
62. Freitag, C. S., and B. I. Eisenstein. 1983. Genetic mapping and transcriptional orientation of the fimD gene. J. Bacteriol. 156:1052–1058.[PubMed]
63. Froehlich, B., L. Husmann, J. Caron, and J. R. Scott. 1994. Regulation of rns, a positive regulatory factor for pili of enterotoxigenic Escherichia coli. J. Bacteriol. 176:5385–5392.[PubMed]
64. Gallegos, M. T., R. Schleif, A. Bairoch, K. Hofmann, and J. L. Ramos. 1997. Arac/XylS family of transcriptional regulators. Microbiol. Mol. Biol. Rev. 61:393–410.[PubMed]
65. Gally, D. L., J. A. Bogan, B. I. Eisenstein, and I. C. Blomfield. 1993. Environmental regulation of the fim switch controlling type 1 fimbrial phase variation in Escherichia coli K-12: effects of temperature and media. J. Bacteriol. 175:6186–6193.[PubMed]
66. Gally, D. L., J. Leathart, and I. C. Blomfield. 1996. Interaction of FimB and FimE with the fim switch that controls the phase variation of type 1 fimbriae in Escherichia coli K-12. Mol. Microbiol. 21:725–738.[PubMed] [CrossRef]
67. Gally, D. L., T. J. Rucker, and I. C. Blomfield. 1994. The leucine-responsive regulatory protein binds to the fim switch to control phase variation of type 1 fimbrial expression in Escherichia coli K-12. J. Bacteriol. 176:5665–5672.[PubMed]
68. Gerstel, U., A. Kolb, and U. Romling. 2006. Regulatory components at the csgD promoter: additional roles for OmpR and integration host factor and role of the 5’ untranslated region. FEMS Microbiol. Lett. 261:109–117.[PubMed] [CrossRef]
69. Gerstel, U., C. Park, and U. Romling. 2003. Complex regulation of csgD promoter activity by global regulatory proteins. Mol. Microbiol. 49:639–654.[PubMed] [CrossRef]
70. Gerstel, U., and U. Romling. 2003. The csgD promoter, a control unit for biofilm formation in Salmonella typhimurium. Res. Microbiol. 154:659–667.[PubMed] [CrossRef]
71. Goransson, M., K. Forsman, and B. E. Uhlin. 1988. Functional and structural homology among regulatory cistrons of pili-adhesin determinants in Escherichia coli. Mol. Gen. Genet. 212:412–417. [CrossRef]
72. Goransson, M., P. Forsman, P. Nilsson, and B. E. Uhlin. 1989. Upstream activating sequences that are shared by two divergently transcribed operons mediate cAMP-CRP regulation of pilus-adhesin in Escherichia coli. Mol. Microbiol. 3:1557–1565.[PubMed] [CrossRef]
73. Guillier, M., and S. Gottesman. 2006. Remodelling of the Escherichia coli outer membrane by two small regulatory RNAs. Mol. Microbiol. 59:231–247.[PubMed] [CrossRef]
74. Gunther, N. W., IV, V. Lockatell, D. E. Johnson, and H. L. Mobley. 2001. In vivo dynamics of type 1 fimbria regulation in uropathogenic Escherichia coli during experimental urinary tract infection. Infect. Immun. 69:2838–2846.[PubMed] [CrossRef]
75. Guyer, D. M., J. S. Kao, and H. L. Mobley. 1998. Genomic analysis of a pathogenicity island in uropathogenic Escherichia coli CFT073: distribution of homologous sequences among isolates from patients with pyelonephritis, cystitis, and catheter-associated bacteriuria and from fecal samples. Infect. Immun. 66:4411–4417.[PubMed]
76. Hammar, M., A. Arnqvist, Z. Bian, A. Olsen, and S. Normark. 1995. Expression of two csg operons is required for production of fibronectin- and Congo red-binding curli polymers in Escherichia coli K-12. Mol. Microbiol. 18:661–670.[PubMed] [CrossRef]
77. Harel, J., F. Daigle, C. Forget, M. C. Tessier, C. Crost, and C. Martin. 2000. Phase variation of F165(1) (Prs-like) fimbriae from Escherichia coli causing septicaemia in animals. Can. J. Microbiol. 46:1101–1107.[PubMed] [CrossRef]
78. Hernday, A., B. Braaten, and D. Low. 2004. The intricate workings of a bacterial epigenetic switch. Adv. Exp. Med. Biol. 547:83–89.[PubMed]
79. Hernday, A., M. Krabbe, B. Braaten, and D. Low. 2002. Self-perpetuating epigenetic pili switches in bacteria. Proc. Natl. Acad. Sci. USA 99(Suppl. 4):16470–16476. [CrossRef]
80. Hernday, A. D., B. A. Braaten, G. Broitman-Maduro, P. Engelberts, and D. A. Low. 2004. Regulation of the pap epigenetic switch by CpxAR: phosphorylated CpxR inhibits transition to the phase ON state by competition with Lrp. Mol. Cell 16:537–547.[PubMed] [CrossRef]
81. Hernday, A. D., B. A. Braaten, and D. A. Low. 2003. The mechanism by which DNA adenine methylase and PapI activate the pap epigenetic switch. Mol. Cell 12:947–957.[PubMed] [CrossRef]
82. Higgins, C. F., C. J. Dorman, D. A. Stirling, L. Waddell, I. R. Booth, G. May, and E. Bremer. 1988. A physiological role for DNA supercoiling in the osmotic regulation of gene expression in S. typhimurium and E. coli. Cell 52:569–584.[PubMed] [CrossRef]
83. Hinde, P., P. Deighan, and C. J. Dorman. 2005. Characterization of the detachable Rho-dependent transcription terminator of the fimE gene in Escherichia coli K-12. J. Bacteriol. 187:8256–8266.[PubMed] [CrossRef]
84. Holden, N., C. Cotterill, and D. Gally. 2000. Examination of regulatory cross-talk between the decay accelerating factor-binding fimbrial/afimbrial adhesins and type I fimbriae. Adv. Exp. Med. Biol. 485:143–150.[PubMed] [CrossRef]
85. Holden, N. J., and D. L. Gally. 2004. Switches, cross-talk and memory in Escherichia coli adherence. J. Med. Microbiol. 53:585–593.[PubMed] [CrossRef]
86. Holden, N. J., M. Totsika, E. Mahler, A. J. Roe, K. Catherwood, K. Lindner, U. Dobrindt, and D. L. Gally. 2006. Demonstration of regulatory cross-talk between P fimbriae and type 1 fimbriae in uropathogenic Escherichia coli. Microbiology 152:1143–1153.[PubMed] [CrossRef]
87. Holden, N. J., B. E. Uhlin, and D. L. Gally. 2001. PapB paralogues and their effect on the phase variation of type 1 fimbriae in Escherichia coli. Mol. Microbiol. 42:319–330.[PubMed] [CrossRef]
88. Honarvar, S., B. K. Choi, and D. M. Schifferli. 2003. Phase variation of the 987P-like CS18 fimbriae of human enterotoxigenic Escherichia coli is regulated by site-specific recombinases. Mol. Microbiol. 48:157–171.[PubMed] [CrossRef]
89. Huisman, T. T., D. Bakker, P. Klaasen, and F. K. de Graaf. 1994. Leucine-responsive regulatory protein, IS1 insertions, and the negative regulator FaeA control the expression of the fae (K88) operon in Escherichia coli. Mol. Microbiol. 11:525–536.[PubMed] [CrossRef]
90. Huisman, T. T., and F. K. de Graaf. 1995. Negative control of fae (K88) expression by the ‘global’ regulator Lrp is modulated by the ‘local’ regulator FaeA and affected by DNA methylation. Mol. Microbiol. 16:943–953.[PubMed] [CrossRef]
91. Hultgren, S. J., W. R. Schwan, A. J. Schaeffer, and J. L. Duncan. 1986. Regulation of production of type 1 pili among urinary tract isolates of Escherichia coli. Infect. Immun. 54:613–620.[PubMed]
92. Humphries, A., S. Deridder, and A. J. Baumler. 2005. Salmonella enterica serotype Typhimurium fimbrial proteins serve as antigens during infection of mice. Infect. Immun. 73:5329–5338.[PubMed] [CrossRef]
93. Humphries, A. D., M. Raffatellu, S. Winter, E. H. Weening, R. A. Kingsley, R. Droleskey, S. Zhang, J. Figueiredo, S. Khare, J. Nunes, L. G. Adams, R. M. Tsolis, and A. J. Baumler. 2003. The use of flow cytometry to detect expression of subunits encoded by 11 Salmonella enterica serotype Typhimurium fimbrial operons. Mol. Microbiol. 48:1357–1376.[PubMed] [CrossRef]
94. Hung, D. L., T. L. Raivio, C. H. Jones, T. J. Silhavy, and S. J. Hultgren. 2001. Cpx signaling pathway monitors biogenesis and affects assembly and expression of P pili. EMBO J. 20:1508–1518.[PubMed] [CrossRef]
95. Isaacson, R. E. 1983. Regulation of expression of Escherichia coli pilus K99. Infect. Immun. 40:633–639.[PubMed]
96. Jones, A. M., A. Goodwill, and T. Elliott. 2006. Limited role for the DsrA and RprA regulatory RNAs in rpoS regulation in Salmonella enterica. J. Bacteriol. 188:5077–5088.[PubMed] [CrossRef]
97. Joyce, S. A., and C. J. Dorman. 2002. A Rho-dependent phase-variable transcription terminator controls expression of the FimE recombinase in Escherichia coli. Mol. Microbiol. 45:1107–1117.[PubMed] [CrossRef]
98. Jubelin, G., A. Vianney, C. Beloin, J. M. Ghigo, J. C. Lazzaroni, P. Lejeune, and C. Dorel. 2005. CpxR/OmpR interplay regulates curli gene expression in response to osmolarity in Escherichia coli. J. Bacteriol. 187:2038–2049.[PubMed] [CrossRef]
99. Kader, A., R. Simm, U. Gerstel, M. Morr, and U. Romling. 2006. Hierarchical involvement of various GGDEF domain proteins in rdar morphotype development of Salmonella enterica serovar Typhimurium. Mol. Microbiol. 60:602–616.[PubMed] [CrossRef]
100. Kalivoda, K. A., S. M. Steenbergen, E. R. Vimr, and J. Plumbridge. 2003. Regulation of sialic acid catabolism by the DNA binding protein NanR in Escherichia coli. J. Bacteriol. 185:4806–4815.[PubMed] [CrossRef]
101. Kaltenbach, L., B. Braaten, J. Tucker, M. Krabbe, and D. Low. 1998. Use of a two-color genetic screen to identify a domain of the global regulator Lrp that is specifically required for pap phase variation. J. Bacteriol. 180:1224–1231.[PubMed]
102. Kaltenbach, L. S., B. A. Braaten, and D. A. Low. 1995. Specific binding of PapI to Lrp-pap DNA complexes. J. Bacteriol. 177:6449–6455.[PubMed]
103. Kao, J. S., D. M. Stucker, J. W. Warren, and H. L. Mobley. 1997. Pathogenicity island sequences of pyelonephritogenic Escherichia coli CFT073 are associated with virulent uropathogenic strains. Infect. Immun. 65:2812–2820.[PubMed]
104. Kawula, T. H., and P. E. Orndorff. 1991. Rapid site-specific DNA inversion in Escherichia coli mutants lacking the histonelike protein H-NS. J. Bacteriol. 173:4116–4123.[PubMed]
105. Kikuchi, T., Y. Mizunoe, A. Takade, S. Naito, and S. Yoshida. 2005. Curli fibers are required for development of biofilm architecture in Escherichia coli K-12 and enhance bacterial adherence to human uroepithelial cells. Microbiol. Immunol. 49:875–884.[PubMed]
106. Kingsley, R. A., E. H. Weening, A. M. Keestra, and A. J. Baumler. 2002. Population heterogeneity of Salmonella enterica serotype Typhimurium resulting from phase variation of the lpf operon in vitro and in vivo. J. Bacteriol. 184:2352–2359.[PubMed] [CrossRef]
107. Klaasen, P., and F. K. de Graaf. 1990. Characterization of FapR, a positive regulator of expression of the 987P operon in enterotoxigenic Escherichia coli. Mol. Microbiol. 4:1779–1783.[PubMed] [CrossRef]
108. Klemm, P. 1986. Two regulatory fim genes, fimB and fimE, control the phase variation of type 1 fimbriae in Escherichia coli. EMBO J. 5:1389–1393.[PubMed]
109. Koo, J. T., J. Choe, and S. L. Moseley. 2004. HrpA, a DEAH-box RNA helicase, is involved in mRNA processing of a fimbrial operon in Escherichia coli. Mol. Microbiol. 52:1813–1826.[PubMed] [CrossRef]
110. Kulasekara, H. D., and I. C. Blomfield. 1999. The molecular basis for the specificity of fimE in the phase variation of type 1 fimbriae of Escherichia coli K-12. Mol. Microbiol. 31:1171–1181.[PubMed] [CrossRef]
111. Lahooti, M., P. L. Roesch, and I. C. Blomfield. 2005. Modulation of the sensitivity of FimB recombination to branched-chain amino acids and alanine in Escherichia coli K-12. J. Bacteriol. 187:6273–6280.[PubMed] [CrossRef]
112. Landgraf, J. R., J. Wu, and J. M. Calvo. 1996. Effects of nutrition and growth rate on Lrp levels in Escherichia coli. J. Bacteriol. 178:6930–6936.[PubMed]
113. Latasa, C., A. Roux, A. Toledo-Arana, J. M. Ghigo, C. Gamazo, J. R. Penades, and I. Lasa. 2005. BapA, a large secreted protein required for biofilm formation and host colonization of Salmonella enterica serovar Enteritidis. Mol. Microbiol. 58:1322–1339.[PubMed]
114. Leathart, J. B., and D. L. Gally. 1998. Regulation of type 1 fimbrial expression in uropathogenic Escherichia coli: heterogeneity of expression through sequence changes in the fim switch region. Mol. Microbiol. 28:371–381.[PubMed] [CrossRef]
115. Lee, Y. M., P. A. DiGiuseppe, T. J. Silhavy, and S. J. Hultgren. 2004. P pilus assembly motif necessary for activation of the CpxRA pathway by PapE in Escherichia coli. J. Bacteriol. 186:4326–4337.[PubMed] [CrossRef]
116. Li, X., D. A. Rasko, C. V. Lockatell, D. E. Johnson, and H. L. Mobley. 2001. Repression of bacterial motility by a novel fimbrial gene product. EMBO J. 20:4854–4862.[PubMed] [CrossRef]
117. Lim, J. K., N. W. Gunther IV, H. Zhao, D. E. Johnson, S. K. Keay, and H. L. Mobley. 1998. In vivo phase variation of Escherichia coli type 1 fimbrial genes in women with urinary tract infection. Infect. Immun. 66:3303–3310.[PubMed]
118. Loomis, W. P., J. T. Koo, T. P. Cheung, and S. L. Moseley. 2001. A tripeptide sequence within the nascent DaaP protein is required for mRNA processing of a fimbrial operon in Escherichia coli. Mol. Microbiol. 39:693–707.[PubMed] [CrossRef]
119. Loomis, W. P., and S. L. Moseley. 1998. Translational control of mRNA processing in the F1845 fimbrial operon of Escherichia coli. Mol. Microbiol. 30:843–853.[PubMed] [CrossRef]
120. Low, D., E. N. Robinson, Jr., Z. A. McGee, and S. Falkow. 1987. The frequency of expression of pyelonephritis-associated pili is under regulatory control. Mol. Microbiol. 1:335–346.[PubMed] [CrossRef]
121. Maiti, S. N., L. DesGroseillers, J. M. Fairbrother, and J. Harel. 1994. Analysis of genes coding for the major and minor fimbrial subunits of the Prs-like fimbriae F165(1) of porcine septicemic Escherichia coli strain 4787. Microb. Pathog. 16:15–25.[PubMed] [CrossRef]
122. Martin, C. 1996. The clp (CS31A) operon is negatively controlled by Lrp, ClpB, and L-alanine at the transcriptional level. Mol. Microbiol. 21:281–292.[PubMed] [CrossRef]
123. Martin, R. G., and J. L. Rosner. 2001. The AraC transcriptional activators. Curr. Opin. Microbiol. 4:132–137.[PubMed] [CrossRef]
124. McClain, M. S., I. C. Blomfield, K. J. Eberhardt, and B. I. Eisenstein. 1993. Inversion-independent phase variation of type 1 fimbriae in Escherichia coli. J. Bacteriol. 175:4335–4344.[PubMed]
125. McClain, M. S., I. C. Blomfield, and B. I. Eisenstein. 1991. Roles of fimB and fimE in site-specific DNA inversion associated with phase variation of type 1 fimbriae in Escherichia coli. J. Bacteriol. 173:5308–5314.[PubMed]
126. McClelland, M., K. E. Sanderson, J. Spieth, S. W. Clifton, P. Latreille, L. Courtney, S. Porwollik, J. Ali, M. Dante, F. Du, S. Hou, D. Layman, S. Leonard, C. Nguyen, K. Scott, A. Holmes, N. Grewal, E. Mulvaney, E. Ryan, H. Sun, L. Florea, W. Miller, T. Stoneking, M. Nhan, R. Waterston, and R. K. Wilson. 2001. Complete genome sequence of Salmonella enterica serovar Typhimurium LT2. Nature 413:852–856.[PubMed] [CrossRef]
127. Mobley, H. L., K. G. Jarvis, J. P. Elwood, D. I. Whittle, C. V. Lockatell, R. G. Russell, D. E. Johnson, M. S. Donnenberg, and J. W. Warren. 1993. Isogenic P-fimbrial deletion mutants of pyelonephritogenic Escherichia coli: the role of alpha Gal(1–4) beta Gal binding in virulence of a wild-type strain. Mol. Microbiol. 10:143–155.[PubMed] [CrossRef]
128. Morschhauser, J., V. Vetter, T. Korhonen, B. E. Uhlin, and J. Hacker. 1993. Regulation and binding properties of S fimbriae cloned from E. coli strains causing urinary tract infection and meningitis. Zentbl. Bakteriol. 278:165–176.
129. Muller, K. H., S. K. Collinson, T. J. Trust, and W. W. Kay. 1991. Type 1 fimbriae of Salmonella enteritidis. J. Bacteriol. 173:4765–4772.[PubMed]
130. Munson, G. P., L. G. Holcomb, and J. R. Scott. 2001. Novel group of virulence activators within the AraC family that are not restricted to upstream binding sites. Infect. Immun. 69:186–193.[PubMed] [CrossRef]
131. Munson, G. P., and J. R. Scott. 1999. Binding site recognition by Rns, a virulence regulator in the AraC family. J. Bacteriol. 181:2110–2117.[PubMed]
132. Munson, G. P., and J. R. Scott. 2000. Rns, a virulence regulator within the AraC family, requires binding sites upstream and downstream of its own promoter to function as an activator. Mol. Microbiol. 36:1391–1402.[PubMed] [CrossRef]
133. Murphree, D., B. Froehlich, and J. R. Scott. 1997. Transcriptional control of genes encoding CS1 pili: negative regulation by a silencer and positive regulation by Rns. J. Bacteriol. 179:5736–5743.[PubMed]
134. Nagy, B., and P. Z. Fekete. 1999. Enterotoxigenic Escherichia coli (ETEC) in farm animals. Vet. Res. 30:259–284.[PubMed]
135. Nataro, J. P., D. Yikang, D. Yingkang, and K. Walker. 1994. AggR, a transcriptional activator of aggregative adherence fimbria I expression in enteroaggregative Escherichia coli. J. Bacteriol. 176:4691–4699.[PubMed]
136. Nicholson, B., and D. Low. 2000. DNA methylation-dependent regulation of pef expression in Salmonella typhimurium. Mol. Microbiol. 35:728–742.[PubMed] [CrossRef]
137. Nilsson, P., S. Naureckiene, and B. E. Uhlin. 1996. Mutations affecting mRNA processing and fimbrial biogenesis in the Escherichia coli pap operon. J. Bacteriol. 178:683–690.[PubMed]
138. Nilsson, P., and B. E. Uhlin. 1991. Differential decay of a polycistronic Escherichia coli transcript is initiated by RNaseE-dependent endonucleolytic processing. Mol. Microbiol. 5:1791–1799.[PubMed] [CrossRef]
139. Normark, S., D. Lark, R. Hull, M. Norgren, M. Baga, P. O'Hanley, G. Schoolnik, and S. Falkow. 1983. Genetics of digalactoside-binding adhesion from a uropathogenic Escherichia coli strain. Infect. Immun. 41:942–949.[PubMed]
140. Norris, T. L., R. A. Kingsley, and A. J. Bumler. 1998. Expression and transcriptional control of the Salmonella typhimurium Ipf fimbrial operon by phase variation. Mol. Microbiol. 29:311–320.[PubMed] [CrossRef]
141. Nou, X., B. Braaten, L. Kaltenbach, and D. A. Low. 1995. Differential binding of Lrp to two sets of pap DNA binding sites mediated by Pap I regulates Pap phase variation in Escherichia coli. EMBO J. 14:5785–5797.[PubMed]
142. Nou, X., B. Skinner, B. Braaten, L. Blyn, D. Hirsch, and D. Low. 1993. Regulation of pyelonephritis-associated pili phase-variation in Escherichia coli: binding of the PapI and the Lrp regulatory proteins is controlled by DNA methylation. Mol. Microbiol. 7:545–553.[PubMed] [CrossRef]
143. Nowicki, J., M. Rhen, V. Vaisanen-Rhen, A. Pere, and T. K. Korhonen. 1984. Immunofluorescence study of fimbrial phase variation in Escherichia coli KS71. J. Bacteriol. 160:691–695.[PubMed]
144. O'Gara, J. P., and C. J. Dorman. 2000. Effects of local transcription and H-NS on inversion of the fim switch of Escherichia coli. Mol. Microbiol. 36:457–466.[PubMed] [CrossRef]
145. Olsen, A., A. Arnqvist, M. Hammar, S. Sukupolvi, and S. Normark. 1993. The RpoS sigma factor relieves H-NS-mediated transcriptional repression of csgA, the subunit gene of fibronectin-binding curli in Escherichia coli. Mol. Microbiol. 7:523–536.[PubMed] [CrossRef]
146. Olsen, A., A. Jonsson, and S. Normark. 1989. Fibronectin binding mediated by a novel class of surface organelles on Escherichia coli. Nature 338:652–655.[PubMed] [CrossRef]
147. Olsen, P. B., and P. Klemm. 1994. Localization of promoters in the fim gene cluster and the effect of H-NS on the transcription of fimB and fimE. FEMS Microbiol. Lett. 116:95–100.[PubMed] [CrossRef]
148. Olsen, P. B., M. A. Schembri, D. L. Gally, and P. Klemm. 1998. Differential temperature modulation by H-NS of the fimB and fimE recombinase genes which control the orientation of the type 1 fimbrial phase switch. FEMS Microbiol. Lett. 162:17–23.[PubMed] [CrossRef]
149. Pallesen, L., O. Madsen, and P. Klemm. 1989. Regulation of the phase switch controlling expression of type 1 fimbriae in Escherichia coli. Mol. Microbiol. 3:925–931.[PubMed] [CrossRef]
150. Paul, B. J., M. B. Berkmen, and R. L. Gourse. 2005. DksA potentiates direct activation of amino acid promoters by ppGpp. Proc. Natl. Acad. Sci. USA 102:7823–7828.[PubMed] [CrossRef]
151. Peterson, S. N., and N. O. Reich. 2006. GATC flanking sequences regulate Dam activity: evidence for how Dam specificity may influence pap expression. J. Mol. Biol. 355:459–472.[PubMed] [CrossRef]
152. Plumbridge, J. 2001. DNA binding sites for the Mlc and NagC proteins: regulation of nagE, encoding the N-acetylglucosamine-specific transporter in Escherichia coli. Nucleic Acids Res. 29:506–514.[PubMed] [CrossRef]
153. Pratt, L. A., and T. J. Silhavy. 1998. Crl stimulates RpoS activity during stationary phase. Mol. Microbiol. 29:1225–1236.[PubMed] [CrossRef]
154. Prigent-Combaret, C., E. Brombacher, O. Vidal, A. Ambert, P. Lejeune, P. Landini, and C. Dorel. 2001. Complex regulatory network controls initial adhesion and biofilm formation in Escherichia coli via regulation of the csgD gene. J. Bacteriol. 183:7213–7223.[PubMed] [CrossRef]
155. Prigent-Combaret, C., G. Prensier, T. T. Le Thi, O. Vidal, P. Lejeune, and C. Dorel. 2000. Developmental pathway for biofilm formation in curli-producing Escherichia coli strains: role of flagella, curli and colanic acid. Environ. Microbiol. 2:450–464.[PubMed] [CrossRef]
156. Provence, D. L., and R. Curtiss III. 1992. Role of crl in avian pathogenic Escherichia coli: a knockout mutation of crl does not affect hemagglutination activity, fibronectin binding, or curli production. Infect. Immun. 60:4460–4467.[PubMed]
157. Repoila, F., N. Majdalani, and S. Gottesman. 2003. Small non-coding RNAs, co-ordinators of adaptation processes in Escherichia coli: the RpoS paradigm. Mol. Microbiol. 48:855–861.[PubMed] [CrossRef]
158. Ritter, A., D. L. Gally, P. B. Olsen, U. Dobrindt, A. Friedrich, P. Klemm, and J. Hacker. 1997. The Pai-associated leuX specific tRNA5(Leu) affects type 1 fimbriation in pathogenic Escherichia coli by control of FimB recombinase expression. Mol. Microbiol. 25:871–882.[PubMed] [CrossRef]
159. Robbe-Saule, V., V. Jaumouille, M. C. Prevost, S. Guadagnini, C. Talhouarne, H. Mathout, A. Kolb, and F. Norel. 2006. Crl activates transcription initiation of RpoS-regulated genes involved in the multicellular behavior of Salmonella enterica serovar Typhimurium. J. Bacteriol. 188:3983–3994.[PubMed] [CrossRef]
160. Robinson, L. S., E. M. Ashman, S. J. Hultgren, and M. R. Chapman. 2006. Secretion of curli fibre subunits is mediated by the outer membrane-localized CsgG protein. Mol. Microbiol. 59:870–881.[PubMed] [CrossRef]
161. Roesch, P. L., and I. C. Blomfield. 1998. Leucine alters the interaction of the leucine-responsive regulatory protein (Lrp) with the fim switch to stimulate site-specific recombination in Escherichia coli. Mol. Microbiol. 27:751–761.[PubMed] [CrossRef]
162. Romling, U. 2005. Characterization of the rdar morphotype, a multicellular behaviour in Enterobacteriaceae. Cell. Mol. Life Sci. 62:1234–1246.[PubMed] [CrossRef]
163. Romling, U., and D. Amikam. 2006. Cyclic di-GMP as a second messenger. Curr. Opin. Microbiol. 9:218–228.[PubMed] [CrossRef]
164. Romling, U., Z. Bian, M. Hammar, W. D. Sierralta, and S. Normark. 1998. Curli fibers are highly conserved between Salmonella typhimurium and Escherichia coli with respect to operon structure and regulation. J. Bacteriol. 180:722–731.[PubMed]
165. Romling, U., M. Rohde, A. Olsen, S. Normark, and J. Reinkoster. 2000. AgfD, the checkpoint of multicellular and aggregative behaviour in Salmonella typhimurium regulates at least two independent pathways. Mol. Microbiol. 36:10–23.[PubMed] [CrossRef]
166. Romling, U., W. D. Sierralta, K. Eriksson, and S. Normark. 1998. Multicellular and aggregative behaviour of Salmonella typhimurium strains is controlled by mutations in the agfD promoter. Mol. Microbiol. 28:249–264.[PubMed] [CrossRef]
167. Roosendaal, B., J. Damoiseaux, W. Jordi, and F. K. de Graaf. 1989. Transcriptional organization of the DNA region controlling expression of the K99 gene cluster. Mol. Gen. Genet. 215:250–256.[PubMed] [CrossRef]
168. Roosendaal, E., M. Boots, and F. K. de Graaf. 1987. Two novel genes, fanA and fanB, involved in the biogenesis of K99 fimbriae. Nucleic Acids Res. 15:5973–5984.[PubMed] [CrossRef]
169. Ruiz, N., and T. J. Silhavy. 2005. Sensing external stress: watchdogs of the Escherichia coli cell envelope. Curr. Opin. Microbiol. 8:122–126.[PubMed] [CrossRef]
170. Sakellaris, H., N. K. Hannink, K. Rajakumar, D. Bulach, M. Hunt, C. Sasakawa, and B. Adler. 2000. Curli loci of Shigella spp. Infect. Immun. 68:3780–3783.[PubMed] [CrossRef]
171. Schembri, M. A., P. B. Olsen, and P. Klemm. 1998. Orientation-dependent enhancement by H-NS of the activity of the type 1 fimbrial phase switch promoter in Escherichia coli. Mol. Gen. Genet. 259:336–344.[PubMed] [CrossRef]
172. Schmoll, T., J. Morschhauser, M. Ott, B. Ludwig, I. van Die, and J. Hacker. 1990. Complete genetic organization and functional aspects of the Escherichia coli S fimbrial adhesin determinant: nucleotide sequence of the genes sfa B, C, D, E, F. Microb. Pathog. 9:331–343.[PubMed] [CrossRef]
173. Schwan, W. R., J. L. Lee, F. A. Lenard, B. T. Matthews, and M. T. Beck. 2002. Osmolarity and pH growth conditions regulate fim gene transcription and type 1 pilus expression in uropathogenic Escherichia coli. Infect. Immun. 70:1391–1402.[PubMed] [CrossRef]
174. Schwan, W. R., H. S. Seifert, and J. L. Duncan. 1994. Analysis of the fimB promoter region involved in type 1 pilus phase variation in Escherichia coli. Mol. Gen. Genet. 242:623–630.[PubMed] [CrossRef]
175. Schwan, W. R., H. S. Seifert, and J. L. Duncan. 1992. Growth conditions mediate differential transcription of fim genes involved in phase variation of type 1 pili. J. Bacteriol. 174:2367–2375.[PubMed]
176. Smith, S. G., and C. J. Dorman. 1999. Functional analysis of the FimE integrase of Escherichia coli K-12: isolation of mutant derivatives with altered DNA inversion preferences. Mol. Microbiol. 34:965–979.[PubMed] [CrossRef]
177. Smyth, C. J. 1982. Two mannose-resistant haemagglutinins on enterotoxigenic Escherichia coli of serotype O6:K15:H16 or H isolated from travellers’ and infantile diarrhoea. J. Gen. Microbiol. 128:2081–2096.[PubMed]
178. Snyder, J. A., B. J. Haugen, C. V. Lockatell, N. Maroncle, E. C. Hagan, D. E. Johnson, R. A. Welch, and H. L. Mobley. 2005. Coordinate expression of fimbriae in uropathogenic Escherichia coli. Infect. Immun. 73:7588–7596.[PubMed] [CrossRef]
179. Snyder, J. A., A. L. Lloyd, C. V. Lockatell, D. E. Johnson, and H. L. Mobley. 2006. Role of phase variation of type 1 fimbriae in a uropathogenic Escherichia coli cystitis isolate during urinary tract infection. Infect. Immun. 74:1387–1393.[PubMed] [CrossRef]
180. Snyder, W. B., L. J. Davis, P. N. Danese, C. L. Cosma, and T. J. Silhavy. 1995. Overproduction of NlpE, a new outer membrane lipoprotein, suppresses the toxicity of periplasmic LacZ by activation of the Cpx signal transduction pathway. J. Bacteriol. 177:4216–4223.[PubMed]
181. Sohanpal, B. K., S. El-Labany, M. Lahooti, J. A. Plumbridge, and I. C. Blomfield. 2004. Integrated regulatory responses of fimB to N-acetylneuraminic (sialic) acid and GlcNAc in Escherichia coli K-12. Proc. Natl. Acad. Sci. USA 101:16322–16327.[PubMed] [CrossRef]
182. Sohanpal, B. K., H. D. Kulasekara, A. Bonnen, and I. C. Blomfield. 2001. Orientational control of fimE expression in Escherichia coli. Mol. Microbiol. 42:483–494.[PubMed] [CrossRef]
183. Sohanpal, B. K., S. Friar, J. Roobol, J. A. Plumbridge, and I. C. Blomfield. 2007. Multiple co-regulatory elements and IHF are necessary for the control of fimB expression in response to sialic acid and N-acetylglucosamine in Escherichia coli K-12. Mol. Microbiol. 63:1223–1236.[PubMed] [CrossRef]
184. Spears, P. A., D. Schauer, and P. E. Orndorff. 1986. Metastable regulation of type 1 piliation in Escherichia coli and isolation and characterization of a phenotypically stable mutant. J. Bacteriol. 168:179–185.[PubMed]
185. Stentebjerg-Olesen, B., T. Chakraborty, and P. Klemm. 2000. FimE-catalyzed off-to-on inversion of the type 1 fimbrial phase switch and insertion sequence recruitment in an Escherichia coli K-12 fimB strain. FEMS Microbiol. Lett. 182:319–325.[PubMed] [CrossRef]
186. Swenson, D. L., K. J. Kim, E. W. Six, and S. Clegg. 1994. The gene fimU affects expression of Salmonella typhimurium type 1 fimbriae and is related to the Escherichia coli tRNA gene argU. Mol. Gen. Genet. 244:216–218.[PubMed] [CrossRef]
187. Tinker, J. K., and S. Clegg. 2000. Characterization of FimY as a coactivator of type 1 fimbrial expression in Salmonella enterica serovar Typhimurium. Infect. Immun. 68:3305–3313.[PubMed] [CrossRef]
188. Tinker, J. K., and S. Clegg. 2001. Control of FimY translation and type 1 fimbrial production by the arginine tRNA encoded by fimU in Salmonella enterica serovar Typhimurium. Mol. Microbiol. 40:757–768.[PubMed] [CrossRef]
189. Tinker, J. K., L. S. Hancox, and S. Clegg. 2001. FimW is a negative regulator affecting type 1 fimbrial expression in Salmonella enterica serovar Typhimurium. J. Bacteriol. 183:435–442.[PubMed] [CrossRef]
190. Uhlich, G. A., P. H. Cooke, and E. B. Solomon. 2006. Analyses of the red-dry-rough phenotype of an Escherichia coli O157:H7 strain and its role in biofilm formation and resistance to antibacterial agents. Appl. Environ. Microbiol. 72:2564–2572.[PubMed] [CrossRef]
191. Uhlich, G. A., J. E. Keen, and R. O. Elder. 2002. Variations in the csgD promoter of Escherichia coli O157:H7 associated with increased virulence in mice and increased invasion of HEp-2 cells. Infect. Immun. 70:395–399.[PubMed] [CrossRef]
192. van der Woude, M., B. Braaten, and D. Low. 1996. Epigenetic phase variation of the pap operon in Escherichia coli. Trends Microbiol. 4:5–9.[PubMed] [CrossRef]
193. van der Woude, M., W. B. Hale, and D. A. Low. 1998. Formation of DNA methylation patterns: nonmethylated GATC sequences in gut and pap operons. J. Bacteriol. 180:5913–5920.[PubMed]
194. van der Woude, M. W., B. A. Braaten, and D. A. Low. 1992. Evidence for global regulatory control of pilus expression in Escherichia coli by Lrp and DNA methylation: model building based on analysis of pap. Mol. Microbiol. 6:2429–2435.[PubMed]
195. van der Woude, M. W., F. K. de Graaf, and H. W. van Verseveld. 1989. Production of the fimbrial adhesin 987P by enterotoxigenic Escherichia coli during growth under controlled conditions in a chemostat. J. Gen. Microbiol. 135:3421–3429.[PubMed]
196. van der Woude, M. W., L. S. Kaltenbach, and D. A. Low. 1995. Leucine-responsive regulatory protein plays dual roles as both an activator and a repressor of the Escherichia coli pap fimbrial operon. Mol. Microbiol. 17:303–312.[PubMed] [CrossRef]
197. van der Woude, M. W., and D. A. Low. 1994. Leucine-responsive regulatory protein and deoxyadenosine methylase control the phase variation and expression of the sfa and daa pili operons in Escherichia coli. Mol. Microbiol. 11:605–618.[PubMed] [CrossRef]
198. Verdonck, F., E. Cox, and B. M. Goddeeris. 2004. F4 fimbriae expressed by porcine enterotoxigenic Escherichia coli, an example of an eccentric fimbrial system? J. Mol. Microbiol. Biotechnol. 7:155–169.[PubMed] [CrossRef]
199. Vianney, A., G. Jubelin, S. Renault, C. Dorel, P. Lejeune, and J. C. Lazzaroni. 2005. Escherichia coli tol and rcs genes participate in the complex network affecting curli synthesis. Microbiology 151:2487–2497.[PubMed] [CrossRef]
200. Viboud, G. I., G. Jonson, E. Dean-Nystrom, and A. M. Svennerholm. 1996. The structural gene encoding human enterotoxigenic Escherichia coli PCFO20 is homologous to that for porcine 987P. Infect. Immun. 64:1233–1239.[PubMed]
201. Vidal, O., R. Longin, C. Prigent-Combaret, C. Dorel, M. Hooreman, and P. Lejeune. 1998. Isolation of an Escherichia coli K-12 mutant strain able to form biofilms on inert surfaces: involvement of a new ompR allele that increases curli expression. J. Bacteriol. 180:2442–2449.[PubMed]
202. Welch, R. A., V. Burland, G. Plunkett III, P. Redford, P. Roesch, D. Rasko, E. L. Buckles, S. R. Liou, A. Boutin, J. Hackett, D. Stroud, G. F. Mayhew, D. J. Rose, S. Zhou, D. C. Schwartz, N. T. Perna, H. L. Mobley, M. S. Donnenberg, and F. R. Blattner. 2002. Extensive mosaic structure revealed by the complete genome sequence of uropathogenic Escherichia coli. Proc. Natl. Acad. Sci. USA 99:17020–17024.[PubMed] [CrossRef]
203. Weyand, N. J., B. A. Braaten, M. van der Woude, J. Tucker, and D. A. Low. 2001. The essential role of the promoter-proximal subunit of CAP in pap phase variation: Lrp- and helical phase-dependent activation of papBA transcription by CAP from −215. Mol. Microbiol. 39:1504–1522.[PubMed] [CrossRef]
204. Weyand, N. J., and D. A. Low. 2000. Regulation of Pap phase variation. Lrp is sufficient for the establishment of the phase off pap DNA methylation pattern and repression of pap transcription in vitro. J. Biol. Chem. 275:3192–3200.[PubMed] [CrossRef]
205. White-Ziegler, C. A., M. L. Angus Hill, B. A. Braaten, M. W. van der Woude, and D. A. Low. 1998. Thermoregulation of Escherichia coli pap transcription: H-NS is a temperature-dependent DNA methylation blocking factor. Mol. Microbiol. 28:1121–1137.[PubMed] [CrossRef]
206. White-Ziegler, C. A., A. M. Black, S. H. Eliades, S. Young, and K. Porter. 2002. The N-acetyltransferase RimJ responds to environmental stimuli to repress pap fimbrial transcription in Escherichia coli. J. Bacteriol. 184:4334–4342.[PubMed] [CrossRef]
207. White-Ziegler, C. A., L. B. Blyn, B. A. Braaten, and D. A. Low. 1990. Identification of an Escherichia coli genetic locus involved in thermoregulation of the pap operon. J. Bacteriol. 172:1775–1782.[PubMed]
208. White-Ziegler, C. A., and D. A. Low. 1992. Thermoregulation of the pap operon: evidence for the involvement of RimJ, the N-terminal acetylase of ribosomal protein S5. J. Bacteriol. 174:7003–7012.[PubMed]
209. White-Ziegler, C. A., A. Villapakkam, K. Ronaszeki, and S. Young. 2000. H-NS controls pap and daa fimbrial transcription in Escherichia coli in response to multiple environmental cues. J. Bacteriol. 182:6391–6400.[PubMed] [CrossRef]
210. Wullt, B., G. Bergsten, H. Connell, P. Rollano, N. Gebratsedik, L. Hang, and C. Svanborg. 2001. P-fimbriae trigger mucosal responses to Escherichia coli in the human urinary tract. Cell. Microbiol. 3:255–264.[PubMed] [CrossRef]
211. Wullt, B., G. Bergsten, H. Connell, P. Rollano, N. Gebretsadik, R. Hull, and C. Svanborg. 2000. P fimbriae enhance the early establishment of Escherichia coli in the human urinary tract. Mol. Microbiol. 38:456–464.[PubMed] [CrossRef]
212. Xia, Y., K. Forsman, J. Jass, and B. E. Uhlin. 1998. Oligomeric interaction of the PapB transcriptional regulator with the upstream activating region of pili adhesin gene promoters in Escherichia coli. Mol. Microbiol. 30:513–523.[PubMed] [CrossRef]
213. Xia, Y., D. Gally, K. Forsman-Semb, and B. E. Uhlin. 2000. Regulatory cross-talk between adhesin operons in Escherichia coli: inhibition of type 1 fimbriae expression by the PapB protein. EMBO J. 19:1450–1457.[PubMed] [CrossRef]
214. Xia, Y., and B. E. Uhlin. 1999. Mutational analysis of the PapB transcriptional regulator in Escherichia coli. Regions important for DNA binding and oligomerization. J. Biol. Chem. 274:19723–19730.[PubMed] [CrossRef]
215. Xie, Y., Y. Yao, V. Kolisnychenko, C. H. Teng, and K. S. Kim. 2006. HbiF regulates type 1 fimbriation independently of FimB and FimE. Infect. Immun. 74:4039–4047.[PubMed] [CrossRef]
216. Yeh, K. S., L. S. Hancox, and S. Clegg. 1995. Construction and characterization of a fimZ mutant of Salmonella typhimurium. J. Bacteriol. 177:6861–6865.[PubMed]
217. Yeh, K. S., J. K. Tinker, and S. Clegg. 2002. FimZ binds the Salmonella typhimurium fimA promoter region and may regulate its own expression with FimY. Microbiol. Immunol. 46:1–10.[PubMed]
218. Zogaj, X., M. Nimtz, M. Rohde, W. Bokranz, and U. Romling. 2001. The multicellular morphotypes of Salmonella typhimurium and Escherichia coli produce cellulose as the second component of the extracellular matrix. Mol. Microbiol. 39:1452–1463.[PubMed] [CrossRef]
Module2.4.2.2
