Characterization of Enterohemorrhagic Escherichia coli Strains Based on Acid Resistance Phenotypes
http://www.100md.com
感染与免疫杂志 2005年第8期
Produce Quality and Safety Laboratory, Henry A. Wallace Beltsville Agricultural Research Center, Agricultural Research Service, USDA, Bldg. 002, 10300 Baltimore Avenue, Beltsville, Maryland 20705-2350
Division of Virulence Assessment, Food and Drug Administration, Laurel, Maryland 20708
ABSTRACT
Acid resistance is perceived to be an important property of enterohemorrhagic Escherichia coli strains, enabling the organisms to survive passage through the acidic environment of the stomach so that they may colonize the mammalian gastrointestinal tract and cause disease. Accordingly, the organism has developed at least three genetically and physiologically distinct acid resistance systems which provide different levels of protection. The glutamate-dependent acid resistance (GDAR) system utilizes extracellular glutamate to protect cells during extreme acid challenges and is believed to provide the highest protection from stomach acidity. In this study, the GDAR system of 82 pathogenic E. coli isolates from 34 countries and 23 states within the United States was examined. Twenty-nine isolates were found to be defective in inducing GDAR under aerobic growth conditions, while five other isolates were defective in GDAR under aerobic, as well as fermentative, growth conditions. We introduced rpoS on a low-copy-number plasmid into 26 isolates and were able to restore GDAR in 20 acid-sensitive isolates under aerobic growth conditions. Four isolates were found to be defective in the newly discovered LuxR-like regulator GadE (formerly YhiE). Defects in other isolates could be due to a mutation(s) in a gene(s) with an as yet undefined role in acid resistance since GadE and/or RpoS could not restore acid resistance. These results show that in addition to mutant alleles of rpoS, mutations in gadE exist in natural populations of pathogenic E. coli. Such mutations most likely alter the infectivity of individual isolates and may play a significant role in determining the infective dose of enterohemorrhagic E. coli.
INTRODUCTION
Shiga toxin (Stx)-producing Escherichia coli strains are a worldwide cause of human disease, with a wide spectrum of symptoms ranging from mild diarrhea to life-threatening hemolytic-uremic syndrome (HUS). Most of these pathogenic strains possess other virulence characteristics such as the ability to cause attaching-and-effacing lesions on mucosal epithelial cells of the large intestine. The Stx-producing family of human disease-associated E. coli is also characterized by its diversity of toxin type (i.e., lysogenic for Stx1-encoding phage or Stx2-encoding phage or with both lysogens) and by its O:H serotype range (21, 29).
In spite of diverse virulence characteristics, one common trait that emerges very clearly is that most of these strains have the ability to withstand gastric acidity (18, 23, 30, 50). In fact, acid tolerance plays a vital role in the survival and virulence of diarrheagenic E. coli strains (8, 36, 38). Sigma factor RpoS plays a significant role in survival as it controls several stress-related genes such as catalase, osmotic shock, and heat shock, as well as acid stress (9, 33, 35). Under aerobic growth conditions, rpoS regulates two of the three acid resistance pathways (13, 17, 24). However, the ability to survive exposure to acid is a complex phenotype, which depends on the growth phase, medium, and species of enteric bacteria (4, 10, 23). Two types of acid resistance pathways have been identified in the aerobically grown stationary-phase cells of E. coli and Shigella flexneri (5, 23, 51). The first acid resistance system is referred to as the glucose-repressible oxidative pathway and protects cells above pH 3.0 (23, 24, 38). The structural components of this acid resistance system (other than RpoS), as well as the mechanisms by which it protects the cells, are still unknown (2, 8). The second system is glutamate-dependent acid resistance (GDAR) and can protect cells below pH 3 (19, 38). GDAR requires glutamate decarboxylase (encoded by paralogous genes gadA and gadB), exogenous glutamate, and an antiporter to release the decarboxylation product -aminobutyric acid (encoded by gadC) (8). Regulation of the GDAR system is extremely complex. Based on results from multiple gene array studies with E. coli, in addition to RpoS and the histone-like protein H-NS, several new regulatory proteins have been implicated to play a role in the expression of gadA and gadBC and acid resistance (26, 28). For example, GadX and GadW belong to the AraC/XylS-like family of transcriptional regulators and have been shown to affect gadA/BC expression and acid resistance of cells grown under rich-medium growth conditions (27, 46, 48), whereas GadE (formerly YhiE) is required for acid resistance in cells grown under either rich- or minimal-medium growth conditions (26). In addition, a small RNA molecule as a regulator of acid response genes has also been predicted (34).
Few studies have directly compared the three acid resistance systems of pathogenic E. coli. For example, 13 of 58 Stx-producing E. coli isolates from various sources contained mutated rpoS and exhibited increased acid sensitivity; however, acid challenge studies were performed without considering the possibility that exposure to different growth conditions might influence the ultimate outcome of the acid challenge (50). Recent analysis of molecular aspects of acid tolerance pathways in E. coli has opened up new strategies capable of dissecting individual acid resistance pathways, as well as identification of additional genes that might influence the acid resistance phenotype. Our aim in this study was to analyze the induction of the GDAR system in various pathogenic E. coli strains and investigate the involvement of rpoS and other genes in acid tolerance. This has led to a refinement of the acid resistance phenotype in enterohemorrhagic, human-associated, pathogenic E. coli.
MATERIALS AND METHODS
Bacterial strains and culture conditions. The strains of E. coli used are listed in Table 1. Culture purity of pathogenic E. coli isolates was checked by examining the catalase activity (31) of at least 96 colonies of each strain. Frozen stocks maintained at –75°C were streaked on Luria-Bertani (LB) agar, and after overnight growth at 37°C, a single colony was inoculated into minimal E medium (49) containing 0.4% glucose at pH 7.0 (EG minimal medium) (24) or into the complex LB growth medium buffered with 100 mM MES (morpholineethanesulfonic acid, pH 5.5). Most diarrheagenic E. coli strains are auxotrophs and require amino acids or vitamins for growth in minimal medium (1, 29). The precise vitamin and amino acid requirements of individual strains were not determined; instead, EG minimal medium was supplemented with 50 μg of yeast extract per ml. Strain 52 is auxotrophic for thiamine, nicotinamide, and riboflavin (1). The strain had the same GDAR phenotype when grown in EG medium supplemented with vitamins or EG medium supplemented with yeast extract (5). Strain MG1655 is not an auxotroph, and the addition of yeast extract to EG medium had no influence on its GDAR phenotype. To obtain stationary-phase cultures, cells were grown in the medium on an orbital shaker (220 rpm, 37°C) for 20 to 22 h to an optical density at 600 nm (OD600) of 3.5 or higher. Strains which were subjected to electroporation with a plasmid (pPS4.4, pIB1, pIB2, or pACYC184) were grown in the medium described above supplemented with chloramphenicol (35 μg ml–1).
For fermentative growth under semiaerobic conditions, cultures were inoculated in brain heart infusion broth with 0.4% glucose (BHIG, 3 ml) in test tubes (13 by 100 mm), which were placed at a 45 ° inclination angle, for 18 to 20 h at 37°C under aeration (shaking at 220 rpm) conditions (10, 23).
Acid challenge and heat shock assays. For the amino acid-dependent acid resistance systems, the cells were diluted directly from the growth medium (1:200) to EG medium (pH 2.0) supplemented with either glutamate (1.5 mM for GDAR or AR2) or arginine (1.5 mM for arginine-dependent acid resistance) and challenged at 37°C for 1 h. EG medium was prewarmed to 37°C, and the pH was adjusted with 6 N HCl (4). Control acid challenge experiments were performed with EG medium (pH 2) without addition of glutamate or arginine. Viable counts were determined after acid challenge by diluting cells in phosphate-buffered saline (50 mM, pH 7.2) and plating immediately on LB agar.
For heat shock assays, stationary-phase cells from LB-MES broth were diluted directly (1:200) into phosphate-buffered saline pre-equilibrated to 58°C. Viable counts were determined at 10-s and 1.5-, 3-, 5-, and 7.5-min intervals by diluting cells in phosphate-buffered saline and plating them immediately on LB agar medium.
Western blot analysis. Cultures were incubated for 18 to 22 h at 37°C to the stationary growth phase (OD600 of 3.5 or higher) in EG or LB medium. Cells of each strain were collected by centrifugation (12,500 x g, 4 min), washed twice in saline, resuspended at 1 OD600 U/ml in 1x loading buffer (50 mM Tris-HCl, pH 6.8, 2% sodium dodecyl sulfate, 10% glycerol, 2.5% -mercaptoethanol, 0.01% Na-azide, 0.1% bromophenol blue), and placed in a boiling water bath for 5 min to make a whole-cell protein sample. Each protein extract was fractionated on sodium dodecyl sulfate-polyacrylamide gradient gels (4 to 20%). After electrophoretic transfer of proteins onto nitrocellulose membranes, the glutamate decarboxylase isozymes were revealed by using a rabbit primary antibody raised against synthetic polypeptide 439EDYKASLKYLSDHPKLQ455, corresponding to the sequence at the C-terminal end of GadA/B (Research Genetics, Huntsville, AL), which was able to detect both isozymes (52.6 kDa). An anti-rabbit antibody raised in a goat and coupled to peroxidase was used as a secondary antibody. To detect RpoS, nitrocellulose membranes were treated with mouse monoclonal antibody (NeoClone Biotechnology International, Madison, WI) raised against the 38-kDa (RpoS) subunit of E. coli RNA polymerase, followed by anti-mouse antibody raised in a goat and coupled to peroxidase as a secondary antibody. A SuperSignal chemiluminescence assay kit (Pierce Biotechnology, Rockford, IL) was used to quantitate the primary antibody, and the signal was captured on preflashed Kodak X-ray film.
Caco-2 cell adherence assays. The human Caco-2 intestinal cell line was obtained from the American Type Culture Collection (ATCC; Manassas, VA). Frozen stock cultures were maintained in GIBCO cell freezing medium held at –140°C. The tissue culture cells were cultivated at 36°C in a 94% air-6%CO2 atmosphere in minimal essential medium supplemented with Earle's salts, 10% fetal bovine serum, 2 mM L-glutamine, 0.1 mM nonessential amino acids, and 1 mM sodium pyruvate. All cell culture media and supplements were obtained from GIBCO (Invitrogen, Gaithersburg, MD).
Adherence assays were performed essentially as described earlier by Kopecko et al. (20). In each well of a 24-well cell culture cluster, approximately 2 x 107 mid-log-phase bacteria (OD650 = 0.3 to 0.4) were added to confluent monolayers of about 5 x 105 epithelial cells per well. Adherence was allowed to occur for 1 h at 37°C in a 94% air-6% CO2 atmosphere. The monolayers were washed three times with Hanks' balanced salt solution and then lysed with 0.1% Triton X-100 in 0.9% saline, and adherent bacteria were enumerated by spread plate count onto Trypticase soy agar (BBL, Becton Dickinson Microbiology Systems, Cockeysville, MD). Adherence ability was expressed as the percentage of the inoculum surviving the washing treatment (i.e., percent recovery). All assays were conducted in quadruplicate and independently repeated at least twice. Results are expressed as an average of the replicate experiments. Recovery percentage data were calculated and analyzed as described previously by using the Student t test and one-factor analysis of variance, followed by Dunn or Student-Newman-Keuls multiple comparisons of means when significant differences (P < 0.05) were found (43).
DNA manipulations and cloning procedures. Standard molecular biology methods were used for chromosomal and plasmid DNA isolation, restriction enzyme digestion and ligation, electroporation, and transformations (37). For cloning of the gadE gene from various acid resistance phenotype group D strains, PCR was performed using a 1:9 mixture of Vent and Taq DNA polymerase enzymes as previously described (12). PCR products were cloned in pGEMTeasy vector (Promega Corporation, Madison, WI). DNA sequencing was performed using double-stranded plasmid DNA templates using SP6 and T7 primers at the Iowa State University Sequencing Facility (Ames, IA) using a PCR-based dideoxynucleotide terminator protocol and an Applied Biosystems automated sequencer.
RESULTS
Classification of strains based on induction of glutamate-dependent acid resistance. A total of 82 strains were analyzed and represented isolates obtained from 34 countries, including isolates obtained from 23 states within the United States of America. First, all E. coli strains were classified into groups based on the induction of GDAR after the strains were grown in either EG minimal medium or LB-MES (pH 5.5) or under fermentative growth conditions (Table 2). Those strains which were able to grow in the EG minimal medium with detectable GDAR levels (survival, 1.0%) were classified as acid resistance phenotype group A. There were a total of 28 strains identified in which GDAR was demonstrated to be inducible under these growth conditions (Table 3). Twenty of the 82 strains were able to induce GDAR when grown in LB-MES (pH 5.5) but not when grown in EG minimal medium and were classified as acid resistance phenotype group B (Table 3). None of the 82 strains survived an acid challenge at pH 2 in the absence of glutamate (survival, <0.001%; data not shown). Other strains (29/82) were able to induce GDAR only under fermentative growth conditions (growth on glucose) and were classified as acid resistance phenotype group C (Table 3). Finally, there were five strains which were unable to induce GDAR under any of the induction growth conditions, and these were classified as acid resistance phenotype group D (Table 3). Arginine-dependent acid resistance, which is induced only under fermentative growth conditions, was measured (Table 2) but was not considered for the classification scheme.
As shown in Table 3, there was no correlation between a single acid resistance phenotypic pattern and a dominant lipopolysaccharide-flagellum (O:H) serotype of the diarrheagenic E. coli strains. For example, acid resistance phenotype group A consisted of 27 strains with 11 strains representative of well-recognized enterohemorrhagic E. coli serotypes, including five O157:H7 strains and one strain each of enterohemorrhagic E. coli serotypes O26:HN, O26:H, O55:H7, and O157:NM and two strains with a serotype designation of O113:H21. Acid resistance phenotype group A also consisted of several strains with traditional enteropathogenic E. coli O serogroups or serotypes, including a single O55:H6 strain, five O111 strains with various flagellar antigens, and a single isolate of serotype O119:H6. Acid resistance phenotype group A also contained one enterotoxigenic strain (O128:H7), one enteroaggregative strain (O44:H18), and seven strains with atypical serotype designations, i.e., O103:HN, O104:H21, O111:H9, O121:NM, O128:H21, O128:NM, and O156:H21. Without being too repetitious, the descriptions of the serotype characteristics for the other three acid resistance phenotypic groups are also shown in Table 3.
Glutamate decarboxylase expression profile and glutamate-dependent acid resistance. Next we examined if a correlation existed between the synthesis of glutamate decarboxylase (encoded by gadA and gadB) and the induction of GDAR across each of the four acid resistance groups (Fig. 1). Both rpoS-dependent (aerobic growth to stationary phase in EG and LB-MES media) and rpoS-independent (fermentative growth to stationary phase in BHIG medium) induction pathways were examined for three strains from individual acid resistance groups. The glutamate decarboxylase isozymes (GadA and GadB, 52.6 kDa) are 98% identical at the amino acid level (40), and their synthesis was monitored by Western blot analysis using an antibody raised against a synthetic peptide (4, 5).
Strains 210, 237, and 245, which belong to acid resistance phenotype group A, were able to induce GDAR under aerobic growth condition (EG medium, lanes 1 to 3, and LB-MES medium, lanes 4 to 6, respectively) and had correspondingly high level of GadAB proteins (Fig. 1A). The GadAB proteins were not detected in acid resistance phenotype group B strains 205, 212, and 234 grown in EG medium; however, the proteins could be detected from cells grown in LB-MES, as well as in BHIG (Fig. 1B). The cells had functional GDAR, which protected them at pH 2. Strains 200, 218, and 219, which belonged to acid resistance phenotype group C, could synthesize GadAB only under fermentative growth conditions in BHIG medium (panel C) and did not survive an acid challenge of pH 2 if grown in EG or LB-MES medium. The GadAB expression pattern of acid resistance phenotype group C strains matched very closely rpoS mutant strains 55 and 65 (Fig. 1, reference strain panel). All five strains belonging to acid resistance phenotype group D were unable to synthesize GadAB in either of the growth media (data not shown).
Influence of functional rpoS on GDAR of acid resistance group C strains. The alternative sigma factor (S) encoded by rpoS is involved in regulation of the GDAR system in stationary, aerobically grown cells. When grown under fermentative growth conditions, cells induced the GDAR system in an rpoS-independent manner. In order to confirm that the defect in GDAR in acid resistance phenotype group C strains is primarily due to erroneous rpoS-mediated regulation of GadAB synthesis, we performed genetic complementation tests. By electroporating pPS4.4, which is a moderately low-copy vector construct carrying a functional rpoS gene, recombinant strains were tested for GDAR. We were able to electroporate 26 of the 29 acid resistance phenotype group C strains (strains 99, 220, and 232 were resistant to chloramphenicol). Complementation of functional rpoS by electroporation restored GDAR under aerobic growth conditions in most of the acid resistance phenotype group C strains, except for strains 100, 104, 226, 244, 252, and 255 (data not shown). From this subgroup, strains 226 and 255 were selected for further analysis (see below).
In comparison to their wild-type parental strains, recombinant strains 107, 230, 218, and 219 carrying pPS4.4 synthesized RpoS, which could be detected by Western blot analysis (Fig. 2A). (The wild-type and corresponding recombinant strains carrying pPS4.4 are referred without and with a -1 suffix, e.g., strains 107 and 107-1, respectively.) Concomitantly, the strains synthesized glutamate decarboxylase and were able to induce GDAR when grown aerobically in LB-MES. In the majority of the acid resistance phenotype group C strains, synthesis of RpoS could not be detected by Western blot analysis. On the other hand, RpoS from strains 226, 247, 255, and 256 could be detected by Western blot analysis (Fig. 2B). The translation levels of the protein were clearly reduced in strains 255 and 256 (Fig. 2B, lanes 1 and 3) but were significantly higher for the other two strains (lanes 5 and 7). In spite of their RpoS levels, the strains were unable to synthesize the GadAB proteins and could not induce GDAR when grown to stationary phase in LB-MES medium. Upon electroporation with pPS4.4, much higher levels of RpoS could be detected in strains 255-1 and 256-1 (Fig. 2B, lanes 1 and 3 versus 2 and 4, respectively), while there was no discernible difference in the RpoS levels of strains 247-1 and 226-1 (Fig. 2B, lanes 5 and 7 versus 6 and 8, respectively). However, only strains 256-1 and 247-1 were able to synthesize GadAB, as well as induce GDAR (Fig. 2B, lanes 2 and 6), while other recombinant strains (226-1 and 255-1) remained defective in GDAR during aerobic growth (Fig. 2B, lanes 4 and 8).
In order to ascertain that pPS4.4 encoded catalytically active RpoS in strains 226 and 255, we compared the temperature tolerances of the wild-type and recombinant strains at 58°C (Fig. 3). The survival patterns of strains 255 and 256 were very similar, and both strains acquired considerable temperature tolerance upon receiving pPS4.4, a phenotype indicative of successful rpoS expression in both strains. Similarly, heat shock analysis of strains 247 and 226 indicated that strain 247-1 gained heat tolerance along with GDAR, while strain 226 was heat tolerant to begin with and remained so after receiving pPS4.4.
Effect of functional gadE on induction of GDAR of acid resistance phenotype group D strains. The inability of the acid resistance phenotype group D strains to induce GDAR even under fermentative growth conditions indicated that the cells were not able to utilize either of the rpoS-independent or rpoS-dependent pathways. The recently discovered LuxR-like regulator GadE (formerly YhiE) is suggested to be required in addition to the rpoS+ genetic background for the aerobic induction of GDAR (26). We analyzed the DNA sequence of the gadE gene from strains 206, 209, 227, 241, and 248 and examined functional complementation by mobilizing plasmid pIB1 (gadE) by electroporation (Table 4). All acid resistance phenotype group D strains, with the exception of 209, had a defect in their gadE sequence; however, only two strains were complemented for GDAR when the gadE gene was mobilized by electroporation. The complementation in strain 248 was fully successful, as gadE restored the aerobic (rpoS-dependent) and fermentative (rpoS-independent) induction pathways. The ability of strain 227 (pIB1) to induce GDAR only under fermentative growth conditions indicated the lack of functional rpoS, and to confirm this possibility, pIB2 (rpoS gadE) was mobilized to strain 227. With both regulators being functional, strain 227(pIB2) was fully restored in GDAR induction (Table 4). Two other strains, 206 and 241, remained defective for GDAR in spite of complementing the defective gadE and/or rpoS gene(s).
Influence of gadE and rpoS on Caco-2 cell adherence properties of acid resistance phenotype group D strains 227 and 248. Recently, Tatsuno et al. (44) reported increased adherence to Caco-2 cells due to disruption of the yhiE (now gadE) and yhiF genes in enterohemorrhagic E. coli O157:H7. We examined strains 227 and 248, which carried a mutation in the gadE gene, for their adherence properties on Caco-2 cells (Fig. 4). Salmonella enterica serovar Typhi strain Ty-2 was used as a positive control, and E. coli HB101 adhesion was considered a baseline reading. Contrary to the findings of Tatsuno et al. (44), strain 248 was not more adherent to Caco-2 cells; in fact, strain 248(pIB1) adhered slightly better than strain 248. Similarly, strain 227 and strain 227(pIB1) both adhered to Caco-2 cells at reduced efficiency and were not significantly different than strain HB101. Since strain 227 had mutations in gadE as well as rpoS, we examined 227(pIB2) and 227(pPS4.4) for cell adhesion in order to gauge the influence of individual regulators. Collectively, both regulators positively influenced adhesion of strain 227; however, the influence of gadE was observed only in the rpoS+ genetic background. The other two acid resistance phenotype group D isolates which carried a mutation in gadE, strains 206 and 241, adhered at 0.47% ± 0.26% and 1.44% ± 0.76% efficiency. Strain 209, with the wild-type gadE sequence, adhered at 3.09% ± 1.37% efficiency.
DISCUSSION
In the most comprehensive single study, over 20% (13/58) of Stx-producing E. coli isolates were reported to carry a mutation in rpoS and had reduced acid tolerance (50). In yet another study, a wide range of acid tolerances among different strains of E. coli O157:H7 and E. coli non-O157:H7 isolates was recorded (7). Although studies like these have contributed substantially to our knowledge of acid tolerance of pathogenic E. coli, a more detailed analysis aimed at dissecting individual acid resistance pathways is needed. Since all available evidence suggests that induction of acid resistance occurs prior to ingestion, we analyzed 82 pathogenic E. coli strains for the ability to induce acid resistance systems under different physiological conditions and particularly analyzed and compared induction of GDAR among different strains.
In the beginning, we classified strains on the basis of the ability to induce GDAR when grown in EG minimal medium. Comparative genome sequence analysis studies have indicated that most pathogenic E. coli strains lack up to 10% of the backbone DNA present in K-12 strains (14). We found that approximately 60% of the strains (acid resistance phenotype group B and group C; 49/82) had an auxotrophic requirement for GDAR induction in EG medium that could not be met with 50 μg ml–1 yeast extract and could induce this acid resistance system only in complex medium such as LB-MES or BHIG. Most strains (94%; 77/82) were able to induce GDAR under either aerobic or fermentative growth conditions (all strains except those in acid resistance phenotype group D). Both regulatory pathways, RpoS dependent and RpoS independent for GDAR induction, were defective in acid resistance phenotype group D strains (6% of strains, 5/82). However, the arginine-dependent acid resistance pathway was operative in all acid resistance phenotype group D strains. Thus, the availability of multiple acid resistance pathways ensures the pathogen's life style and at the same time might explain the discrepancies observed in acid resistance and/or the infective dose among various strains as different acid resistance systems may be required to be operative under commensal or pathogenic lifestyles (35, 36).
We found that 29 out of 82 isolates were defective in the aerobic induction of GDAR, which requires functional rpoS. The general stress resistance of E. coli is controlled by the RpoS sigma factor, but mutations in rpoS are surprisingly common in natural and laboratory strains (11, 22, 25, 50). The questions of whether there is any possibility of a selective advantage in losing rpoS and whether RpoS contributes to fitness under nutrient limitations are being actively investigated by several laboratories (15, 16, 22, 32). We analyzed the strains with a putative defect in rpoS by a physiological and recombinant DNA approach. Out of the 26 acid resistance phenotype group C strains we electroporated with pPS4.4 (rpoS), 20 strains restored their ability to synthesize RpoS and GadAB, as well as the ability to induce GDAR under aerobic growth conditions. However, due to constrains of the Western blot assay technique, only limited information could be drawn from strains 256, 255, 247, and 226, which exhibited various levels of RpoS expression, and their GDAR expression was not uniform once they acquired functional rpoS. The reason for the lack of GDAR expression in strains 247 and 226 does not appear to be weak translation or suboptimal quantities of RpoS (Fig. 2B, lanes 5 and 7, respectively), although the possibility remains that the RpoS protein is dysfunctional.
In order to overcome this limitation of the Western blotting technique, we took advantage of the fact that rpoS also regulates heat tolerance and used the heat tolerance assay to gauge the functionality of native and recombinant RpoS (Fig. 3). In spite of detectable levels of RpoS, strain 247 belonged to acid resistance phenotype group C and was heat sensitive. However, the strain gained GDAR, as well as heat tolerance, upon electroporation with pPS4.4, indicating that RpoS from strain 247 could be catalytically inactive. On the contrary, strain 226 does not appear to be defective in rpoS based on its heat tolerance ability (Fig. 3) and it is likely that strain 226(pPS4.4) could be overexpressing RpoS, creating an imbalance in the cell's heat tolerance. It is likely that wild-type strain 226 could be defective in GDAR due to a lack of a regulatory element(s) yet to be identified for aerobic induction. Further more, strain 255 also appears to carry a defect(s) in another regulatory gene(s) in addition to rpoS since in spite of successful expression of RpoS from pPS4.4 (i.e., complementation for heat tolerance), this strain remained defective in GDAR (Fig. 2B). Recently, a small regulatory RNA (gadY) was identified in E. coli K-12 (MG 1665) whose expression is dependent on rpoS and which is predicted to have a role in the enhanced expression of gadX and downstream acid resistance genes (34). Strains 226 and 255 need to be examined for any mutations in gadY; alternatively, the strains could be used as trap hosts to identify regulatory elements for aerobic induction of the GDAR system. It may be noted that all acid resistance phenotype group C isolates were able to induce GDAR by an rpoS-independent pathway when they were grown under fermentative conditions, indicating a fully functional GDAR system (Table 2).
Four isolates from acid resistance phenotype group D were defective in a newly discovered LuxR-type positive regulator, gadE. However, mobilization of pIB1 (gadE) could restore GDAR in strain 248 while in strain 227 it required fermentative growth, indicating a possible defect in rpoS (Table 4), which was later confirmed by complementation using pIB2 (rpoS gadE). The reasons for the inability to induce GDAR (under either aerobic or fermentative growth conditions) by three of the group D isolates (strains 206, 209, and 241) are not apparent at this time. All group D isolates were examined for the presence of gadBC and gadXW regions by PCR amplification (data not shown). Although we could detect identical-size PCR amplification products on agarose gels with reference to the wild-type strains, the possibility of a point mutation(s) or smaller deletions cannot be ruled out. Alternatively, the strains could be lacking regulators yet to be identified for the rpoS-independent induction of GDAR. The possible involvement of yhiE (gadE) in negatively regulating adhesion to intestinal cells in E. coli O157:H7 (44) prompted us to examine group D strains in Caco-2 cell assays. However, we were unable to detect a negative regulatory role of gadE in adhesion to Caco-2 cells. In fact, we observed a synergistic effect of two regulators, and strain 227(pIB2) was most efficient in adhering to Caco-2 cells. Strains 227 and 248 belonged to serogroups O111:H8 and O145:HNM, respectively, and in addition to the involvement of multiple elements in controlling the adherence of E. coli O157:H7 (45), it appears that regulation of adhesion mechanisms differs among O157 and other serogroups (42).
The rpoS mutations are much more prevalent in natural populations, and certainly mutS and rpoS are located in a highly polymorphic segment of the chromosome (6, 11, 16). In studies in which pathogenic or commensal E. coli isolates were analyzed, several sequence variants of rpoS were reported and a range of phenotypes attributable to altered rpoS function were exhibited (3, 9, 22). Based on batch and chemostat culture studies, it has been proposed that mutant rpoS alleles can confer a survival advantage over wild-type strains (16, 22). The GDAR system provides the highest level of protection, functioning at pH 2 or less, and can be induced in an rpoS-independent manner (2, 8). GDAR may also be very important for food safety, as monosodium glutamate will support acid resistance although it is intended to be a food preservative. In order to extend shelf life, more and more ready-to-eat food is marketed in modified-atmosphere packages (41, 52). It is necessary to determine if such an environment would provide fermentative conditions for induction of GDAR in an rpoS-independent manner.
ACKNOWLEDGMENTS
This study was supported in part by the overseas Industrial Attachment Program of the School of Life Sciences and Chemical Technology, Ngee Ann Polytechnic, Singapore.
We are grateful to John Foster for providing strains and plasmids prior to publication. We thank Ingrid Berlanger, Letitia Bolds, Irving Newman, Frances Trouth, and Nhi Vo for excellent technical assistance.
Dedicated to Irving Newman, who passed away on 5 February 2005.
Permanent address: School of Life Sciences and Chemical Technology, Ngee Ann Polytechnic, 535 Clement Road, Singapore 599489, Singapore.
REFERENCES
1. Arnold, C. N., J. McElhanon, A. Lee, R. Leonhart, and D. A. Siegele. 2001. Global analysis of Escherichia coli gene expression during the acetate-induced acid tolerance response. J. Bacteriol. 183:2178-2186.
2. Audia, J. P., C. C. Webb, and J. W. Foster. 2001. Breaking through the acid barrier: an orchestrated response to proton stress by enteric bacteria. Int. J. Med. Microbiol. 291:97-106.
3. Benito, A., G. Ventoura, M. Casadei, T. Robinson, and B. Mackey. 1999. Variation in resistance of natural isolates of Escherichia coli O157 to high hydrostatic pressure, mild heat, and other stresses. Appl. Environ. Microbiol. 65:1564-1569.
4. Bhagwat, A. A. 2003. Regulation of the glutamate-dependent acid-resistant system of diarrheagenic Escherichia coli strains. FEMS Microbiol. Lett. 227:39-45.
5. Bhagwat, A. A., and M. A. Bhagwat. 2004. Comparative analysis of transcriptional regulatory elements of glutamate-dependent acid-resistance systems of Shigella flexneri and Escherichia coli O157:H7. FEMS Microbiol. Lett. 234:139-147.
6. Brown, E. W., J. E. LeClerc, B. Li, W. L. Payne, and T. A. Cebula. 2001. Phylogenetic evidence for horizontal transfer of mutS alleles among naturally occurring Escherichia coli strains. J. Bacteriol. 183:1631-1644.
7. Brudzinski, L., and M. A. Harrison. 1998. Influence of incubation conditions on survival and acid tolerance response of Escherichia coli O157:H7 and non-O157:H7 isolates exposed to acetic acid. J. Food Prot. 61:542-546.
8. Castanie-Cornet, M. P., T. A. Penfound, D. Smith, J. F. Elliott, and J. W. Foster. 1999. Control of acid resistance in Escherichia coli. J. Bacteriol. 181:3525-3535.
9. Cheville, A. M., K. W. Arnold, C. Buchrieser, C. M. Cheng, and C. W. Kaspar. 1996. rpoS regulation of acid, heat, and salt tolerance in Escherichia coli O157:H7. Appl. Environ. Microbiol. 62:1822-1824.
10. Cui, S., J. Meng, and A. A. Bhagwat. 2001. Availability of glutamate and arginine during acid challenge determines cell density-dependent survival phenotype of Escherichia coli strains. Appl. Environ. Microbiol. 67:4914-4918.
11. Culham, D. E., and J. M. Wood. 2000. An Escherichia coli reference collection group B2- and uropathogen-associated polymorphism in the rpoS-mutS region of the E. coli chromosome. J. Bacteriol. 182:6272-6276.
12. Datsenko, K. A., and B. L. Wanner. 2000. One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc. Natl. Acad. Sci. USA 97:6640-6645.
13. De Biase, D., A. Tramonti, F. Bossa, and P. Visca. 1999. The response to stationary-phase stress conditions in Escherichia coli: role and regulation of the glutamic acid decarboxylase system. Mol. Microbiol. 32:1198-1211.
14. Dobrindt, U., F. Agerer, K. Michaelis, A. Janka, C. Buchrieser, M. Samuelson, C. Svanborg, G. Gottschalk, H. Karch, and J. Hacker. 2003. Analysis of genome plasticity in pathogenic and commensal Escherichia coli isolates by use of DNA arrays. J. Bacteriol. 185:1831-1840.
15. Farrell, M. J., and S. E. Finkel. 2003. The growth advantage in stationary-phase phenotype conferred by rpoS mutations is dependent on the pH and nutrient environment. J. Bacteriol. 185:7044-7052.
16. Ferenci, T. 2003. What is driving the acquisition of mutS and rpoS polymorphisms in Escherichia coli Trends Microbiol. 11:457-461.
17. Foster, J. W. 2000. Microbial responses to acid stress, p. 99-115. In G. Storz and R. Hengge-Aronis (ed.), Bacterial stress responses. ASM Press, Washington D.C.
18. Foster, J. W., and M. Moreno. 1999. Inducible acid tolerance mechanisms in enteric bacteria. Novartis Found. Symp. 221:55-69.
19. Hersh, B. M., F. T. Farooq, D. N. Barstad, D. L. Blankenhorn, and J. L. Slonczewski. 1996. A glutamate-dependent acid resistance gene in Escherichia coli. J. Bacteriol. 178:3978-3981.
20. Huang, X. Z., B. D. Tall, W. Schwan, and D. J. Kopecko. 1998. Physical limits of Salmonella typhi entry into cultured human intestinal epithelial cells. Infect. Immun. 66:2928-2937.
21. Kaper, J. B., J. P. Nataro, and H. L. T. Mobley. 2004. Pathogenic Escherichia coli. Nat. Rev. Microbiol. 2:123-140.
22. King, T., A. Ishihama, A. Kori, and T. Ferenci. 2004. A regulatory trade-off as a source of strain variation in the species Escherichia coli. J. Bacteriol. 186:5614-5620.
23. Lin, J., I. S. Lee, J. Frey, J. L. Slonczewski, and J. W. Foster. 1995. Comparative analysis of extreme acid survival in Salmonella typhimurium, Shigella flexneri, and Escherichia coli. J. Bacteriol. 177:4097-4104.
24. Lin, J., M. P. Smith, K. C. Chapin, H. S. Baik, G. N. Bennett, and J. W. Foster. 1996. Mechanisms of acid resistance in enterohemorrhagic Escherichia coli. Appl. Environ. Microbiol. 62:3094-3100.
25. Lombardo, M., I. Aponyi, and S. M. Rosenberg. 2004. General stress response regulator RpoS in adaptive mutation and amplification in Escherichia coli. Genetics 166:669-680.
26. Ma, Z., S. Gong, H. Richard, D. L. Tucker, T. Conway, and J. W. Foster. 2003. GadE (YhiE) activates glutamate decarboxylase-dependent acid resistance in Escherichia coli K-12. Mol. Microbiol. 49:1309-1320.
27. Ma, Z., H. Richard, D. L. Tucker, T. Conway, and J. W. Foster. 2002. Collaborative regulation of Escherichia coli glutamate-dependent acid resistance by two AraC-like regulators, GadX and GadW (YhiW). J. Bacteriol. 184:7001-7012.
28. Masuda, N., and G. M. Church. 2003. Regulatory network of acid resistance genes in Escherichia coli. Mol. Microbiol. 48:699-712.
29. Meng, J., M. P. Dolye, T. Zhao, and S. Zhao. 2001. Enterohemorrhagic Escherichia coli, p. 193-213. In M. P. Doyle, L. R. Beuchat, and T. J. Montville (ed.), Food microbiology: fundamentals and frontiers, 2nd ed. ASM Press, Washington, D.C.
30. Merrell, D. S., and A. Camilli. 2002. Acid tolerance of gastrointestinal pathogens. Curr. Opin. Microbiol. 5:51-55.
31. Notley-McRobb, L., T. King, and T. Ferenci. 2002. rpoS mutations and loss of general stress resistance in Escherichia coli populations as a consequence of conflict between competing stress responses. J. Bacteriol. 184:806-811.
32. Nystrom, T. 2003. Conditional senescence in bacteria: death of the immortals. Mol. Microbiol. 48:17-23.
33. Nystrom, T. 2004. Stationary-phase physiology. Annu. Rev. Microbiol. 58:161-181.
34. Opdyke, J. A., J.-G. Kang, and G. Storz. 2004. GadY, a small-RNA regulator of acid response genes in Escherichia coli. J. Bacteriol. 186:6698-6705.
35. Price, S. B., C. M. Cheng, C. W. Kaspar, J. C. Wright, F. J. DeGraves, T. A. Penfound, M. P. Castanie-Cornet, and J. W. Foster. 2000. Role of rpoS in acid resistance and fecal shedding of Escherichia coli O157:H7. Appl. Environ. Microbiol. 66:632-637.
36. Price, S. B., J. C. Wright, F. J. DeGraves, M. P. Castanie-Cornet, and J. W. Foster. 2004. Acid resistance systems required for survival of Escherichia coli O157:H7 in the bovine gastrointestinal tract and in apple cider are different. Appl. Environ. Microbiol. 70:4792-4799.
37. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
38. Slonczewski, J. L., and J. W. Foster. 1996. pH-regulated genes and survival at acidic pH, p. 1539-1549. In F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: cellular and molecular biology, 2nd ed., vol. 1. ASM Press, Washington, D.C.
39. Small, P., D. Blankenhorn, D. Welty, E. Zinser, and J. L. Slonczewski. 1994. Acid and base resistance in Escherichia coli and Shigella flexneri: role of rpoS and growth pH. J. Bacteriol. 176:1729-1737.
40. Smith, D. K., T. Kassam, B. Singh, and J. F. Elliott. 1992. Escherichia coli has two homologous glutamate decarboxylase genes that map to distinct loci. J. Bacteriol. 174:5820-5826.
41. Soliva-Fortuny, R. C., and O. Martin-Belloso. 2003. New advances in extending the shelf-life of fresh-cut fruits: a review. Trends Food Sci. Technol. 14:341-353.
42. Stevens, M. P., A. J. Roe, I. Vlisidou, P. M. van Diemen, R. M. La Ragione, A. Best, M. J. Woodward, D. L. Gally, and T. S. Wallis. 2004. Mutation of toxB and a truncated version of the efa-1 gene in Escherichia coli O157:H7 influences the expression and secretion of locus of enterocyte effacement-encoded proteins but not intestinal colonization in calves or sheep. Infect. Immun. 72:5402-5411.
43. Tall, B. D., J. F. La Peyre, J. W. Bier, M. D. Miliotis, D. E. Hanes, M. H. Kothary, D. B. Shah, and M. Faisal. 1999. Perkinsus marinus extracellular protease modulates survival of Vibrio vulnificus in eastern oyster hemocytes. Appl. Env. Microbiol. 65:4261-4263.
44. Tatsuno, I., K. Nagano, K. Taguchi, L. Rong, H. Mori, and C. Sasakawa. 2003. Increased adherence to Caco-2 cells caused by disruption of the yhiE and yhiF genes in enterohemorrhagic Escherichia coli O157:H7. Infect. Immun. 71:2598-2606.
45. Torres, A. G., and J. B. Kaper. 2003. Multiple elements controlling adherence of enterohemorrhagic Escherichia coli O157:H7 to HeLa cells. Infect. Immun. 71:4985-4995.
46. Tramonti, A., P. Visca, M. De Canio, M. Falconi, and D. De Biase. 2002. Functional characterization and regulation of gadX, a gene encoding an AraC/XylS-like transcriptional activator of the Escherichia coli glutamic acid decarboxylase system. J. Bacteriol. 184:2603-2613.
47. Tucker, D. L., N. Tucker, and T. Conway. 2002. Gene expression profiling of the pH response in Escherichia coli. J. Bacteriol. 184:6551-6558.
48. Tucker, D. L., N. Tucker, Z. Ma, J. W. Foster, R. L. Miranda, P. S. Cohen, and T. Conway. 2003. Genes of the GadX-GadW regulon in Escherichia coli. J. Bacteriol. 185:3190-3201.
49. Vogel, H. J., and D. M. Bonner. 1956. Acetylornithinase of Escherichia coli: partial purification and some properties. J. Biol. Chem. 218:97-106.
50. Waterman, S. R., and P. L. C. Small. 1996. Characterization of the acid resistance phenotype and rpoS alleles of Shiga-like toxin-producing Escherichia coli. Infect. Immun. 64:2808-2811.
51. Waterman, S. R., and P. L. C. Small. 2003. Identification of the promoter regions and sigma-S-dependent regulation of the gadA and gadBC genes associated with glutamate-dependent acid resistance in Shigella flexneri. FEMS Microbiol. Lett. 225:155-160.
52. Yuan, J. T. C. 2003. Modified atmosphere packaging for shelf-life extension, p. 215-219. In J. S. Novak, G. M. Sapers, and V. K. Juneja (ed.), Microbial safety of minimally processed foods. CRC Press, Inc., New York, N.Y.(Arvind A. Bhagwat, Lynn C)
Division of Virulence Assessment, Food and Drug Administration, Laurel, Maryland 20708
ABSTRACT
Acid resistance is perceived to be an important property of enterohemorrhagic Escherichia coli strains, enabling the organisms to survive passage through the acidic environment of the stomach so that they may colonize the mammalian gastrointestinal tract and cause disease. Accordingly, the organism has developed at least three genetically and physiologically distinct acid resistance systems which provide different levels of protection. The glutamate-dependent acid resistance (GDAR) system utilizes extracellular glutamate to protect cells during extreme acid challenges and is believed to provide the highest protection from stomach acidity. In this study, the GDAR system of 82 pathogenic E. coli isolates from 34 countries and 23 states within the United States was examined. Twenty-nine isolates were found to be defective in inducing GDAR under aerobic growth conditions, while five other isolates were defective in GDAR under aerobic, as well as fermentative, growth conditions. We introduced rpoS on a low-copy-number plasmid into 26 isolates and were able to restore GDAR in 20 acid-sensitive isolates under aerobic growth conditions. Four isolates were found to be defective in the newly discovered LuxR-like regulator GadE (formerly YhiE). Defects in other isolates could be due to a mutation(s) in a gene(s) with an as yet undefined role in acid resistance since GadE and/or RpoS could not restore acid resistance. These results show that in addition to mutant alleles of rpoS, mutations in gadE exist in natural populations of pathogenic E. coli. Such mutations most likely alter the infectivity of individual isolates and may play a significant role in determining the infective dose of enterohemorrhagic E. coli.
INTRODUCTION
Shiga toxin (Stx)-producing Escherichia coli strains are a worldwide cause of human disease, with a wide spectrum of symptoms ranging from mild diarrhea to life-threatening hemolytic-uremic syndrome (HUS). Most of these pathogenic strains possess other virulence characteristics such as the ability to cause attaching-and-effacing lesions on mucosal epithelial cells of the large intestine. The Stx-producing family of human disease-associated E. coli is also characterized by its diversity of toxin type (i.e., lysogenic for Stx1-encoding phage or Stx2-encoding phage or with both lysogens) and by its O:H serotype range (21, 29).
In spite of diverse virulence characteristics, one common trait that emerges very clearly is that most of these strains have the ability to withstand gastric acidity (18, 23, 30, 50). In fact, acid tolerance plays a vital role in the survival and virulence of diarrheagenic E. coli strains (8, 36, 38). Sigma factor RpoS plays a significant role in survival as it controls several stress-related genes such as catalase, osmotic shock, and heat shock, as well as acid stress (9, 33, 35). Under aerobic growth conditions, rpoS regulates two of the three acid resistance pathways (13, 17, 24). However, the ability to survive exposure to acid is a complex phenotype, which depends on the growth phase, medium, and species of enteric bacteria (4, 10, 23). Two types of acid resistance pathways have been identified in the aerobically grown stationary-phase cells of E. coli and Shigella flexneri (5, 23, 51). The first acid resistance system is referred to as the glucose-repressible oxidative pathway and protects cells above pH 3.0 (23, 24, 38). The structural components of this acid resistance system (other than RpoS), as well as the mechanisms by which it protects the cells, are still unknown (2, 8). The second system is glutamate-dependent acid resistance (GDAR) and can protect cells below pH 3 (19, 38). GDAR requires glutamate decarboxylase (encoded by paralogous genes gadA and gadB), exogenous glutamate, and an antiporter to release the decarboxylation product -aminobutyric acid (encoded by gadC) (8). Regulation of the GDAR system is extremely complex. Based on results from multiple gene array studies with E. coli, in addition to RpoS and the histone-like protein H-NS, several new regulatory proteins have been implicated to play a role in the expression of gadA and gadBC and acid resistance (26, 28). For example, GadX and GadW belong to the AraC/XylS-like family of transcriptional regulators and have been shown to affect gadA/BC expression and acid resistance of cells grown under rich-medium growth conditions (27, 46, 48), whereas GadE (formerly YhiE) is required for acid resistance in cells grown under either rich- or minimal-medium growth conditions (26). In addition, a small RNA molecule as a regulator of acid response genes has also been predicted (34).
Few studies have directly compared the three acid resistance systems of pathogenic E. coli. For example, 13 of 58 Stx-producing E. coli isolates from various sources contained mutated rpoS and exhibited increased acid sensitivity; however, acid challenge studies were performed without considering the possibility that exposure to different growth conditions might influence the ultimate outcome of the acid challenge (50). Recent analysis of molecular aspects of acid tolerance pathways in E. coli has opened up new strategies capable of dissecting individual acid resistance pathways, as well as identification of additional genes that might influence the acid resistance phenotype. Our aim in this study was to analyze the induction of the GDAR system in various pathogenic E. coli strains and investigate the involvement of rpoS and other genes in acid tolerance. This has led to a refinement of the acid resistance phenotype in enterohemorrhagic, human-associated, pathogenic E. coli.
MATERIALS AND METHODS
Bacterial strains and culture conditions. The strains of E. coli used are listed in Table 1. Culture purity of pathogenic E. coli isolates was checked by examining the catalase activity (31) of at least 96 colonies of each strain. Frozen stocks maintained at –75°C were streaked on Luria-Bertani (LB) agar, and after overnight growth at 37°C, a single colony was inoculated into minimal E medium (49) containing 0.4% glucose at pH 7.0 (EG minimal medium) (24) or into the complex LB growth medium buffered with 100 mM MES (morpholineethanesulfonic acid, pH 5.5). Most diarrheagenic E. coli strains are auxotrophs and require amino acids or vitamins for growth in minimal medium (1, 29). The precise vitamin and amino acid requirements of individual strains were not determined; instead, EG minimal medium was supplemented with 50 μg of yeast extract per ml. Strain 52 is auxotrophic for thiamine, nicotinamide, and riboflavin (1). The strain had the same GDAR phenotype when grown in EG medium supplemented with vitamins or EG medium supplemented with yeast extract (5). Strain MG1655 is not an auxotroph, and the addition of yeast extract to EG medium had no influence on its GDAR phenotype. To obtain stationary-phase cultures, cells were grown in the medium on an orbital shaker (220 rpm, 37°C) for 20 to 22 h to an optical density at 600 nm (OD600) of 3.5 or higher. Strains which were subjected to electroporation with a plasmid (pPS4.4, pIB1, pIB2, or pACYC184) were grown in the medium described above supplemented with chloramphenicol (35 μg ml–1).
For fermentative growth under semiaerobic conditions, cultures were inoculated in brain heart infusion broth with 0.4% glucose (BHIG, 3 ml) in test tubes (13 by 100 mm), which were placed at a 45 ° inclination angle, for 18 to 20 h at 37°C under aeration (shaking at 220 rpm) conditions (10, 23).
Acid challenge and heat shock assays. For the amino acid-dependent acid resistance systems, the cells were diluted directly from the growth medium (1:200) to EG medium (pH 2.0) supplemented with either glutamate (1.5 mM for GDAR or AR2) or arginine (1.5 mM for arginine-dependent acid resistance) and challenged at 37°C for 1 h. EG medium was prewarmed to 37°C, and the pH was adjusted with 6 N HCl (4). Control acid challenge experiments were performed with EG medium (pH 2) without addition of glutamate or arginine. Viable counts were determined after acid challenge by diluting cells in phosphate-buffered saline (50 mM, pH 7.2) and plating immediately on LB agar.
For heat shock assays, stationary-phase cells from LB-MES broth were diluted directly (1:200) into phosphate-buffered saline pre-equilibrated to 58°C. Viable counts were determined at 10-s and 1.5-, 3-, 5-, and 7.5-min intervals by diluting cells in phosphate-buffered saline and plating them immediately on LB agar medium.
Western blot analysis. Cultures were incubated for 18 to 22 h at 37°C to the stationary growth phase (OD600 of 3.5 or higher) in EG or LB medium. Cells of each strain were collected by centrifugation (12,500 x g, 4 min), washed twice in saline, resuspended at 1 OD600 U/ml in 1x loading buffer (50 mM Tris-HCl, pH 6.8, 2% sodium dodecyl sulfate, 10% glycerol, 2.5% -mercaptoethanol, 0.01% Na-azide, 0.1% bromophenol blue), and placed in a boiling water bath for 5 min to make a whole-cell protein sample. Each protein extract was fractionated on sodium dodecyl sulfate-polyacrylamide gradient gels (4 to 20%). After electrophoretic transfer of proteins onto nitrocellulose membranes, the glutamate decarboxylase isozymes were revealed by using a rabbit primary antibody raised against synthetic polypeptide 439EDYKASLKYLSDHPKLQ455, corresponding to the sequence at the C-terminal end of GadA/B (Research Genetics, Huntsville, AL), which was able to detect both isozymes (52.6 kDa). An anti-rabbit antibody raised in a goat and coupled to peroxidase was used as a secondary antibody. To detect RpoS, nitrocellulose membranes were treated with mouse monoclonal antibody (NeoClone Biotechnology International, Madison, WI) raised against the 38-kDa (RpoS) subunit of E. coli RNA polymerase, followed by anti-mouse antibody raised in a goat and coupled to peroxidase as a secondary antibody. A SuperSignal chemiluminescence assay kit (Pierce Biotechnology, Rockford, IL) was used to quantitate the primary antibody, and the signal was captured on preflashed Kodak X-ray film.
Caco-2 cell adherence assays. The human Caco-2 intestinal cell line was obtained from the American Type Culture Collection (ATCC; Manassas, VA). Frozen stock cultures were maintained in GIBCO cell freezing medium held at –140°C. The tissue culture cells were cultivated at 36°C in a 94% air-6%CO2 atmosphere in minimal essential medium supplemented with Earle's salts, 10% fetal bovine serum, 2 mM L-glutamine, 0.1 mM nonessential amino acids, and 1 mM sodium pyruvate. All cell culture media and supplements were obtained from GIBCO (Invitrogen, Gaithersburg, MD).
Adherence assays were performed essentially as described earlier by Kopecko et al. (20). In each well of a 24-well cell culture cluster, approximately 2 x 107 mid-log-phase bacteria (OD650 = 0.3 to 0.4) were added to confluent monolayers of about 5 x 105 epithelial cells per well. Adherence was allowed to occur for 1 h at 37°C in a 94% air-6% CO2 atmosphere. The monolayers were washed three times with Hanks' balanced salt solution and then lysed with 0.1% Triton X-100 in 0.9% saline, and adherent bacteria were enumerated by spread plate count onto Trypticase soy agar (BBL, Becton Dickinson Microbiology Systems, Cockeysville, MD). Adherence ability was expressed as the percentage of the inoculum surviving the washing treatment (i.e., percent recovery). All assays were conducted in quadruplicate and independently repeated at least twice. Results are expressed as an average of the replicate experiments. Recovery percentage data were calculated and analyzed as described previously by using the Student t test and one-factor analysis of variance, followed by Dunn or Student-Newman-Keuls multiple comparisons of means when significant differences (P < 0.05) were found (43).
DNA manipulations and cloning procedures. Standard molecular biology methods were used for chromosomal and plasmid DNA isolation, restriction enzyme digestion and ligation, electroporation, and transformations (37). For cloning of the gadE gene from various acid resistance phenotype group D strains, PCR was performed using a 1:9 mixture of Vent and Taq DNA polymerase enzymes as previously described (12). PCR products were cloned in pGEMTeasy vector (Promega Corporation, Madison, WI). DNA sequencing was performed using double-stranded plasmid DNA templates using SP6 and T7 primers at the Iowa State University Sequencing Facility (Ames, IA) using a PCR-based dideoxynucleotide terminator protocol and an Applied Biosystems automated sequencer.
RESULTS
Classification of strains based on induction of glutamate-dependent acid resistance. A total of 82 strains were analyzed and represented isolates obtained from 34 countries, including isolates obtained from 23 states within the United States of America. First, all E. coli strains were classified into groups based on the induction of GDAR after the strains were grown in either EG minimal medium or LB-MES (pH 5.5) or under fermentative growth conditions (Table 2). Those strains which were able to grow in the EG minimal medium with detectable GDAR levels (survival, 1.0%) were classified as acid resistance phenotype group A. There were a total of 28 strains identified in which GDAR was demonstrated to be inducible under these growth conditions (Table 3). Twenty of the 82 strains were able to induce GDAR when grown in LB-MES (pH 5.5) but not when grown in EG minimal medium and were classified as acid resistance phenotype group B (Table 3). None of the 82 strains survived an acid challenge at pH 2 in the absence of glutamate (survival, <0.001%; data not shown). Other strains (29/82) were able to induce GDAR only under fermentative growth conditions (growth on glucose) and were classified as acid resistance phenotype group C (Table 3). Finally, there were five strains which were unable to induce GDAR under any of the induction growth conditions, and these were classified as acid resistance phenotype group D (Table 3). Arginine-dependent acid resistance, which is induced only under fermentative growth conditions, was measured (Table 2) but was not considered for the classification scheme.
As shown in Table 3, there was no correlation between a single acid resistance phenotypic pattern and a dominant lipopolysaccharide-flagellum (O:H) serotype of the diarrheagenic E. coli strains. For example, acid resistance phenotype group A consisted of 27 strains with 11 strains representative of well-recognized enterohemorrhagic E. coli serotypes, including five O157:H7 strains and one strain each of enterohemorrhagic E. coli serotypes O26:HN, O26:H, O55:H7, and O157:NM and two strains with a serotype designation of O113:H21. Acid resistance phenotype group A also consisted of several strains with traditional enteropathogenic E. coli O serogroups or serotypes, including a single O55:H6 strain, five O111 strains with various flagellar antigens, and a single isolate of serotype O119:H6. Acid resistance phenotype group A also contained one enterotoxigenic strain (O128:H7), one enteroaggregative strain (O44:H18), and seven strains with atypical serotype designations, i.e., O103:HN, O104:H21, O111:H9, O121:NM, O128:H21, O128:NM, and O156:H21. Without being too repetitious, the descriptions of the serotype characteristics for the other three acid resistance phenotypic groups are also shown in Table 3.
Glutamate decarboxylase expression profile and glutamate-dependent acid resistance. Next we examined if a correlation existed between the synthesis of glutamate decarboxylase (encoded by gadA and gadB) and the induction of GDAR across each of the four acid resistance groups (Fig. 1). Both rpoS-dependent (aerobic growth to stationary phase in EG and LB-MES media) and rpoS-independent (fermentative growth to stationary phase in BHIG medium) induction pathways were examined for three strains from individual acid resistance groups. The glutamate decarboxylase isozymes (GadA and GadB, 52.6 kDa) are 98% identical at the amino acid level (40), and their synthesis was monitored by Western blot analysis using an antibody raised against a synthetic peptide (4, 5).
Strains 210, 237, and 245, which belong to acid resistance phenotype group A, were able to induce GDAR under aerobic growth condition (EG medium, lanes 1 to 3, and LB-MES medium, lanes 4 to 6, respectively) and had correspondingly high level of GadAB proteins (Fig. 1A). The GadAB proteins were not detected in acid resistance phenotype group B strains 205, 212, and 234 grown in EG medium; however, the proteins could be detected from cells grown in LB-MES, as well as in BHIG (Fig. 1B). The cells had functional GDAR, which protected them at pH 2. Strains 200, 218, and 219, which belonged to acid resistance phenotype group C, could synthesize GadAB only under fermentative growth conditions in BHIG medium (panel C) and did not survive an acid challenge of pH 2 if grown in EG or LB-MES medium. The GadAB expression pattern of acid resistance phenotype group C strains matched very closely rpoS mutant strains 55 and 65 (Fig. 1, reference strain panel). All five strains belonging to acid resistance phenotype group D were unable to synthesize GadAB in either of the growth media (data not shown).
Influence of functional rpoS on GDAR of acid resistance group C strains. The alternative sigma factor (S) encoded by rpoS is involved in regulation of the GDAR system in stationary, aerobically grown cells. When grown under fermentative growth conditions, cells induced the GDAR system in an rpoS-independent manner. In order to confirm that the defect in GDAR in acid resistance phenotype group C strains is primarily due to erroneous rpoS-mediated regulation of GadAB synthesis, we performed genetic complementation tests. By electroporating pPS4.4, which is a moderately low-copy vector construct carrying a functional rpoS gene, recombinant strains were tested for GDAR. We were able to electroporate 26 of the 29 acid resistance phenotype group C strains (strains 99, 220, and 232 were resistant to chloramphenicol). Complementation of functional rpoS by electroporation restored GDAR under aerobic growth conditions in most of the acid resistance phenotype group C strains, except for strains 100, 104, 226, 244, 252, and 255 (data not shown). From this subgroup, strains 226 and 255 were selected for further analysis (see below).
In comparison to their wild-type parental strains, recombinant strains 107, 230, 218, and 219 carrying pPS4.4 synthesized RpoS, which could be detected by Western blot analysis (Fig. 2A). (The wild-type and corresponding recombinant strains carrying pPS4.4 are referred without and with a -1 suffix, e.g., strains 107 and 107-1, respectively.) Concomitantly, the strains synthesized glutamate decarboxylase and were able to induce GDAR when grown aerobically in LB-MES. In the majority of the acid resistance phenotype group C strains, synthesis of RpoS could not be detected by Western blot analysis. On the other hand, RpoS from strains 226, 247, 255, and 256 could be detected by Western blot analysis (Fig. 2B). The translation levels of the protein were clearly reduced in strains 255 and 256 (Fig. 2B, lanes 1 and 3) but were significantly higher for the other two strains (lanes 5 and 7). In spite of their RpoS levels, the strains were unable to synthesize the GadAB proteins and could not induce GDAR when grown to stationary phase in LB-MES medium. Upon electroporation with pPS4.4, much higher levels of RpoS could be detected in strains 255-1 and 256-1 (Fig. 2B, lanes 1 and 3 versus 2 and 4, respectively), while there was no discernible difference in the RpoS levels of strains 247-1 and 226-1 (Fig. 2B, lanes 5 and 7 versus 6 and 8, respectively). However, only strains 256-1 and 247-1 were able to synthesize GadAB, as well as induce GDAR (Fig. 2B, lanes 2 and 6), while other recombinant strains (226-1 and 255-1) remained defective in GDAR during aerobic growth (Fig. 2B, lanes 4 and 8).
In order to ascertain that pPS4.4 encoded catalytically active RpoS in strains 226 and 255, we compared the temperature tolerances of the wild-type and recombinant strains at 58°C (Fig. 3). The survival patterns of strains 255 and 256 were very similar, and both strains acquired considerable temperature tolerance upon receiving pPS4.4, a phenotype indicative of successful rpoS expression in both strains. Similarly, heat shock analysis of strains 247 and 226 indicated that strain 247-1 gained heat tolerance along with GDAR, while strain 226 was heat tolerant to begin with and remained so after receiving pPS4.4.
Effect of functional gadE on induction of GDAR of acid resistance phenotype group D strains. The inability of the acid resistance phenotype group D strains to induce GDAR even under fermentative growth conditions indicated that the cells were not able to utilize either of the rpoS-independent or rpoS-dependent pathways. The recently discovered LuxR-like regulator GadE (formerly YhiE) is suggested to be required in addition to the rpoS+ genetic background for the aerobic induction of GDAR (26). We analyzed the DNA sequence of the gadE gene from strains 206, 209, 227, 241, and 248 and examined functional complementation by mobilizing plasmid pIB1 (gadE) by electroporation (Table 4). All acid resistance phenotype group D strains, with the exception of 209, had a defect in their gadE sequence; however, only two strains were complemented for GDAR when the gadE gene was mobilized by electroporation. The complementation in strain 248 was fully successful, as gadE restored the aerobic (rpoS-dependent) and fermentative (rpoS-independent) induction pathways. The ability of strain 227 (pIB1) to induce GDAR only under fermentative growth conditions indicated the lack of functional rpoS, and to confirm this possibility, pIB2 (rpoS gadE) was mobilized to strain 227. With both regulators being functional, strain 227(pIB2) was fully restored in GDAR induction (Table 4). Two other strains, 206 and 241, remained defective for GDAR in spite of complementing the defective gadE and/or rpoS gene(s).
Influence of gadE and rpoS on Caco-2 cell adherence properties of acid resistance phenotype group D strains 227 and 248. Recently, Tatsuno et al. (44) reported increased adherence to Caco-2 cells due to disruption of the yhiE (now gadE) and yhiF genes in enterohemorrhagic E. coli O157:H7. We examined strains 227 and 248, which carried a mutation in the gadE gene, for their adherence properties on Caco-2 cells (Fig. 4). Salmonella enterica serovar Typhi strain Ty-2 was used as a positive control, and E. coli HB101 adhesion was considered a baseline reading. Contrary to the findings of Tatsuno et al. (44), strain 248 was not more adherent to Caco-2 cells; in fact, strain 248(pIB1) adhered slightly better than strain 248. Similarly, strain 227 and strain 227(pIB1) both adhered to Caco-2 cells at reduced efficiency and were not significantly different than strain HB101. Since strain 227 had mutations in gadE as well as rpoS, we examined 227(pIB2) and 227(pPS4.4) for cell adhesion in order to gauge the influence of individual regulators. Collectively, both regulators positively influenced adhesion of strain 227; however, the influence of gadE was observed only in the rpoS+ genetic background. The other two acid resistance phenotype group D isolates which carried a mutation in gadE, strains 206 and 241, adhered at 0.47% ± 0.26% and 1.44% ± 0.76% efficiency. Strain 209, with the wild-type gadE sequence, adhered at 3.09% ± 1.37% efficiency.
DISCUSSION
In the most comprehensive single study, over 20% (13/58) of Stx-producing E. coli isolates were reported to carry a mutation in rpoS and had reduced acid tolerance (50). In yet another study, a wide range of acid tolerances among different strains of E. coli O157:H7 and E. coli non-O157:H7 isolates was recorded (7). Although studies like these have contributed substantially to our knowledge of acid tolerance of pathogenic E. coli, a more detailed analysis aimed at dissecting individual acid resistance pathways is needed. Since all available evidence suggests that induction of acid resistance occurs prior to ingestion, we analyzed 82 pathogenic E. coli strains for the ability to induce acid resistance systems under different physiological conditions and particularly analyzed and compared induction of GDAR among different strains.
In the beginning, we classified strains on the basis of the ability to induce GDAR when grown in EG minimal medium. Comparative genome sequence analysis studies have indicated that most pathogenic E. coli strains lack up to 10% of the backbone DNA present in K-12 strains (14). We found that approximately 60% of the strains (acid resistance phenotype group B and group C; 49/82) had an auxotrophic requirement for GDAR induction in EG medium that could not be met with 50 μg ml–1 yeast extract and could induce this acid resistance system only in complex medium such as LB-MES or BHIG. Most strains (94%; 77/82) were able to induce GDAR under either aerobic or fermentative growth conditions (all strains except those in acid resistance phenotype group D). Both regulatory pathways, RpoS dependent and RpoS independent for GDAR induction, were defective in acid resistance phenotype group D strains (6% of strains, 5/82). However, the arginine-dependent acid resistance pathway was operative in all acid resistance phenotype group D strains. Thus, the availability of multiple acid resistance pathways ensures the pathogen's life style and at the same time might explain the discrepancies observed in acid resistance and/or the infective dose among various strains as different acid resistance systems may be required to be operative under commensal or pathogenic lifestyles (35, 36).
We found that 29 out of 82 isolates were defective in the aerobic induction of GDAR, which requires functional rpoS. The general stress resistance of E. coli is controlled by the RpoS sigma factor, but mutations in rpoS are surprisingly common in natural and laboratory strains (11, 22, 25, 50). The questions of whether there is any possibility of a selective advantage in losing rpoS and whether RpoS contributes to fitness under nutrient limitations are being actively investigated by several laboratories (15, 16, 22, 32). We analyzed the strains with a putative defect in rpoS by a physiological and recombinant DNA approach. Out of the 26 acid resistance phenotype group C strains we electroporated with pPS4.4 (rpoS), 20 strains restored their ability to synthesize RpoS and GadAB, as well as the ability to induce GDAR under aerobic growth conditions. However, due to constrains of the Western blot assay technique, only limited information could be drawn from strains 256, 255, 247, and 226, which exhibited various levels of RpoS expression, and their GDAR expression was not uniform once they acquired functional rpoS. The reason for the lack of GDAR expression in strains 247 and 226 does not appear to be weak translation or suboptimal quantities of RpoS (Fig. 2B, lanes 5 and 7, respectively), although the possibility remains that the RpoS protein is dysfunctional.
In order to overcome this limitation of the Western blotting technique, we took advantage of the fact that rpoS also regulates heat tolerance and used the heat tolerance assay to gauge the functionality of native and recombinant RpoS (Fig. 3). In spite of detectable levels of RpoS, strain 247 belonged to acid resistance phenotype group C and was heat sensitive. However, the strain gained GDAR, as well as heat tolerance, upon electroporation with pPS4.4, indicating that RpoS from strain 247 could be catalytically inactive. On the contrary, strain 226 does not appear to be defective in rpoS based on its heat tolerance ability (Fig. 3) and it is likely that strain 226(pPS4.4) could be overexpressing RpoS, creating an imbalance in the cell's heat tolerance. It is likely that wild-type strain 226 could be defective in GDAR due to a lack of a regulatory element(s) yet to be identified for aerobic induction. Further more, strain 255 also appears to carry a defect(s) in another regulatory gene(s) in addition to rpoS since in spite of successful expression of RpoS from pPS4.4 (i.e., complementation for heat tolerance), this strain remained defective in GDAR (Fig. 2B). Recently, a small regulatory RNA (gadY) was identified in E. coli K-12 (MG 1665) whose expression is dependent on rpoS and which is predicted to have a role in the enhanced expression of gadX and downstream acid resistance genes (34). Strains 226 and 255 need to be examined for any mutations in gadY; alternatively, the strains could be used as trap hosts to identify regulatory elements for aerobic induction of the GDAR system. It may be noted that all acid resistance phenotype group C isolates were able to induce GDAR by an rpoS-independent pathway when they were grown under fermentative conditions, indicating a fully functional GDAR system (Table 2).
Four isolates from acid resistance phenotype group D were defective in a newly discovered LuxR-type positive regulator, gadE. However, mobilization of pIB1 (gadE) could restore GDAR in strain 248 while in strain 227 it required fermentative growth, indicating a possible defect in rpoS (Table 4), which was later confirmed by complementation using pIB2 (rpoS gadE). The reasons for the inability to induce GDAR (under either aerobic or fermentative growth conditions) by three of the group D isolates (strains 206, 209, and 241) are not apparent at this time. All group D isolates were examined for the presence of gadBC and gadXW regions by PCR amplification (data not shown). Although we could detect identical-size PCR amplification products on agarose gels with reference to the wild-type strains, the possibility of a point mutation(s) or smaller deletions cannot be ruled out. Alternatively, the strains could be lacking regulators yet to be identified for the rpoS-independent induction of GDAR. The possible involvement of yhiE (gadE) in negatively regulating adhesion to intestinal cells in E. coli O157:H7 (44) prompted us to examine group D strains in Caco-2 cell assays. However, we were unable to detect a negative regulatory role of gadE in adhesion to Caco-2 cells. In fact, we observed a synergistic effect of two regulators, and strain 227(pIB2) was most efficient in adhering to Caco-2 cells. Strains 227 and 248 belonged to serogroups O111:H8 and O145:HNM, respectively, and in addition to the involvement of multiple elements in controlling the adherence of E. coli O157:H7 (45), it appears that regulation of adhesion mechanisms differs among O157 and other serogroups (42).
The rpoS mutations are much more prevalent in natural populations, and certainly mutS and rpoS are located in a highly polymorphic segment of the chromosome (6, 11, 16). In studies in which pathogenic or commensal E. coli isolates were analyzed, several sequence variants of rpoS were reported and a range of phenotypes attributable to altered rpoS function were exhibited (3, 9, 22). Based on batch and chemostat culture studies, it has been proposed that mutant rpoS alleles can confer a survival advantage over wild-type strains (16, 22). The GDAR system provides the highest level of protection, functioning at pH 2 or less, and can be induced in an rpoS-independent manner (2, 8). GDAR may also be very important for food safety, as monosodium glutamate will support acid resistance although it is intended to be a food preservative. In order to extend shelf life, more and more ready-to-eat food is marketed in modified-atmosphere packages (41, 52). It is necessary to determine if such an environment would provide fermentative conditions for induction of GDAR in an rpoS-independent manner.
ACKNOWLEDGMENTS
This study was supported in part by the overseas Industrial Attachment Program of the School of Life Sciences and Chemical Technology, Ngee Ann Polytechnic, Singapore.
We are grateful to John Foster for providing strains and plasmids prior to publication. We thank Ingrid Berlanger, Letitia Bolds, Irving Newman, Frances Trouth, and Nhi Vo for excellent technical assistance.
Dedicated to Irving Newman, who passed away on 5 February 2005.
Permanent address: School of Life Sciences and Chemical Technology, Ngee Ann Polytechnic, 535 Clement Road, Singapore 599489, Singapore.
REFERENCES
1. Arnold, C. N., J. McElhanon, A. Lee, R. Leonhart, and D. A. Siegele. 2001. Global analysis of Escherichia coli gene expression during the acetate-induced acid tolerance response. J. Bacteriol. 183:2178-2186.
2. Audia, J. P., C. C. Webb, and J. W. Foster. 2001. Breaking through the acid barrier: an orchestrated response to proton stress by enteric bacteria. Int. J. Med. Microbiol. 291:97-106.
3. Benito, A., G. Ventoura, M. Casadei, T. Robinson, and B. Mackey. 1999. Variation in resistance of natural isolates of Escherichia coli O157 to high hydrostatic pressure, mild heat, and other stresses. Appl. Environ. Microbiol. 65:1564-1569.
4. Bhagwat, A. A. 2003. Regulation of the glutamate-dependent acid-resistant system of diarrheagenic Escherichia coli strains. FEMS Microbiol. Lett. 227:39-45.
5. Bhagwat, A. A., and M. A. Bhagwat. 2004. Comparative analysis of transcriptional regulatory elements of glutamate-dependent acid-resistance systems of Shigella flexneri and Escherichia coli O157:H7. FEMS Microbiol. Lett. 234:139-147.
6. Brown, E. W., J. E. LeClerc, B. Li, W. L. Payne, and T. A. Cebula. 2001. Phylogenetic evidence for horizontal transfer of mutS alleles among naturally occurring Escherichia coli strains. J. Bacteriol. 183:1631-1644.
7. Brudzinski, L., and M. A. Harrison. 1998. Influence of incubation conditions on survival and acid tolerance response of Escherichia coli O157:H7 and non-O157:H7 isolates exposed to acetic acid. J. Food Prot. 61:542-546.
8. Castanie-Cornet, M. P., T. A. Penfound, D. Smith, J. F. Elliott, and J. W. Foster. 1999. Control of acid resistance in Escherichia coli. J. Bacteriol. 181:3525-3535.
9. Cheville, A. M., K. W. Arnold, C. Buchrieser, C. M. Cheng, and C. W. Kaspar. 1996. rpoS regulation of acid, heat, and salt tolerance in Escherichia coli O157:H7. Appl. Environ. Microbiol. 62:1822-1824.
10. Cui, S., J. Meng, and A. A. Bhagwat. 2001. Availability of glutamate and arginine during acid challenge determines cell density-dependent survival phenotype of Escherichia coli strains. Appl. Environ. Microbiol. 67:4914-4918.
11. Culham, D. E., and J. M. Wood. 2000. An Escherichia coli reference collection group B2- and uropathogen-associated polymorphism in the rpoS-mutS region of the E. coli chromosome. J. Bacteriol. 182:6272-6276.
12. Datsenko, K. A., and B. L. Wanner. 2000. One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc. Natl. Acad. Sci. USA 97:6640-6645.
13. De Biase, D., A. Tramonti, F. Bossa, and P. Visca. 1999. The response to stationary-phase stress conditions in Escherichia coli: role and regulation of the glutamic acid decarboxylase system. Mol. Microbiol. 32:1198-1211.
14. Dobrindt, U., F. Agerer, K. Michaelis, A. Janka, C. Buchrieser, M. Samuelson, C. Svanborg, G. Gottschalk, H. Karch, and J. Hacker. 2003. Analysis of genome plasticity in pathogenic and commensal Escherichia coli isolates by use of DNA arrays. J. Bacteriol. 185:1831-1840.
15. Farrell, M. J., and S. E. Finkel. 2003. The growth advantage in stationary-phase phenotype conferred by rpoS mutations is dependent on the pH and nutrient environment. J. Bacteriol. 185:7044-7052.
16. Ferenci, T. 2003. What is driving the acquisition of mutS and rpoS polymorphisms in Escherichia coli Trends Microbiol. 11:457-461.
17. Foster, J. W. 2000. Microbial responses to acid stress, p. 99-115. In G. Storz and R. Hengge-Aronis (ed.), Bacterial stress responses. ASM Press, Washington D.C.
18. Foster, J. W., and M. Moreno. 1999. Inducible acid tolerance mechanisms in enteric bacteria. Novartis Found. Symp. 221:55-69.
19. Hersh, B. M., F. T. Farooq, D. N. Barstad, D. L. Blankenhorn, and J. L. Slonczewski. 1996. A glutamate-dependent acid resistance gene in Escherichia coli. J. Bacteriol. 178:3978-3981.
20. Huang, X. Z., B. D. Tall, W. Schwan, and D. J. Kopecko. 1998. Physical limits of Salmonella typhi entry into cultured human intestinal epithelial cells. Infect. Immun. 66:2928-2937.
21. Kaper, J. B., J. P. Nataro, and H. L. T. Mobley. 2004. Pathogenic Escherichia coli. Nat. Rev. Microbiol. 2:123-140.
22. King, T., A. Ishihama, A. Kori, and T. Ferenci. 2004. A regulatory trade-off as a source of strain variation in the species Escherichia coli. J. Bacteriol. 186:5614-5620.
23. Lin, J., I. S. Lee, J. Frey, J. L. Slonczewski, and J. W. Foster. 1995. Comparative analysis of extreme acid survival in Salmonella typhimurium, Shigella flexneri, and Escherichia coli. J. Bacteriol. 177:4097-4104.
24. Lin, J., M. P. Smith, K. C. Chapin, H. S. Baik, G. N. Bennett, and J. W. Foster. 1996. Mechanisms of acid resistance in enterohemorrhagic Escherichia coli. Appl. Environ. Microbiol. 62:3094-3100.
25. Lombardo, M., I. Aponyi, and S. M. Rosenberg. 2004. General stress response regulator RpoS in adaptive mutation and amplification in Escherichia coli. Genetics 166:669-680.
26. Ma, Z., S. Gong, H. Richard, D. L. Tucker, T. Conway, and J. W. Foster. 2003. GadE (YhiE) activates glutamate decarboxylase-dependent acid resistance in Escherichia coli K-12. Mol. Microbiol. 49:1309-1320.
27. Ma, Z., H. Richard, D. L. Tucker, T. Conway, and J. W. Foster. 2002. Collaborative regulation of Escherichia coli glutamate-dependent acid resistance by two AraC-like regulators, GadX and GadW (YhiW). J. Bacteriol. 184:7001-7012.
28. Masuda, N., and G. M. Church. 2003. Regulatory network of acid resistance genes in Escherichia coli. Mol. Microbiol. 48:699-712.
29. Meng, J., M. P. Dolye, T. Zhao, and S. Zhao. 2001. Enterohemorrhagic Escherichia coli, p. 193-213. In M. P. Doyle, L. R. Beuchat, and T. J. Montville (ed.), Food microbiology: fundamentals and frontiers, 2nd ed. ASM Press, Washington, D.C.
30. Merrell, D. S., and A. Camilli. 2002. Acid tolerance of gastrointestinal pathogens. Curr. Opin. Microbiol. 5:51-55.
31. Notley-McRobb, L., T. King, and T. Ferenci. 2002. rpoS mutations and loss of general stress resistance in Escherichia coli populations as a consequence of conflict between competing stress responses. J. Bacteriol. 184:806-811.
32. Nystrom, T. 2003. Conditional senescence in bacteria: death of the immortals. Mol. Microbiol. 48:17-23.
33. Nystrom, T. 2004. Stationary-phase physiology. Annu. Rev. Microbiol. 58:161-181.
34. Opdyke, J. A., J.-G. Kang, and G. Storz. 2004. GadY, a small-RNA regulator of acid response genes in Escherichia coli. J. Bacteriol. 186:6698-6705.
35. Price, S. B., C. M. Cheng, C. W. Kaspar, J. C. Wright, F. J. DeGraves, T. A. Penfound, M. P. Castanie-Cornet, and J. W. Foster. 2000. Role of rpoS in acid resistance and fecal shedding of Escherichia coli O157:H7. Appl. Environ. Microbiol. 66:632-637.
36. Price, S. B., J. C. Wright, F. J. DeGraves, M. P. Castanie-Cornet, and J. W. Foster. 2004. Acid resistance systems required for survival of Escherichia coli O157:H7 in the bovine gastrointestinal tract and in apple cider are different. Appl. Environ. Microbiol. 70:4792-4799.
37. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
38. Slonczewski, J. L., and J. W. Foster. 1996. pH-regulated genes and survival at acidic pH, p. 1539-1549. In F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: cellular and molecular biology, 2nd ed., vol. 1. ASM Press, Washington, D.C.
39. Small, P., D. Blankenhorn, D. Welty, E. Zinser, and J. L. Slonczewski. 1994. Acid and base resistance in Escherichia coli and Shigella flexneri: role of rpoS and growth pH. J. Bacteriol. 176:1729-1737.
40. Smith, D. K., T. Kassam, B. Singh, and J. F. Elliott. 1992. Escherichia coli has two homologous glutamate decarboxylase genes that map to distinct loci. J. Bacteriol. 174:5820-5826.
41. Soliva-Fortuny, R. C., and O. Martin-Belloso. 2003. New advances in extending the shelf-life of fresh-cut fruits: a review. Trends Food Sci. Technol. 14:341-353.
42. Stevens, M. P., A. J. Roe, I. Vlisidou, P. M. van Diemen, R. M. La Ragione, A. Best, M. J. Woodward, D. L. Gally, and T. S. Wallis. 2004. Mutation of toxB and a truncated version of the efa-1 gene in Escherichia coli O157:H7 influences the expression and secretion of locus of enterocyte effacement-encoded proteins but not intestinal colonization in calves or sheep. Infect. Immun. 72:5402-5411.
43. Tall, B. D., J. F. La Peyre, J. W. Bier, M. D. Miliotis, D. E. Hanes, M. H. Kothary, D. B. Shah, and M. Faisal. 1999. Perkinsus marinus extracellular protease modulates survival of Vibrio vulnificus in eastern oyster hemocytes. Appl. Env. Microbiol. 65:4261-4263.
44. Tatsuno, I., K. Nagano, K. Taguchi, L. Rong, H. Mori, and C. Sasakawa. 2003. Increased adherence to Caco-2 cells caused by disruption of the yhiE and yhiF genes in enterohemorrhagic Escherichia coli O157:H7. Infect. Immun. 71:2598-2606.
45. Torres, A. G., and J. B. Kaper. 2003. Multiple elements controlling adherence of enterohemorrhagic Escherichia coli O157:H7 to HeLa cells. Infect. Immun. 71:4985-4995.
46. Tramonti, A., P. Visca, M. De Canio, M. Falconi, and D. De Biase. 2002. Functional characterization and regulation of gadX, a gene encoding an AraC/XylS-like transcriptional activator of the Escherichia coli glutamic acid decarboxylase system. J. Bacteriol. 184:2603-2613.
47. Tucker, D. L., N. Tucker, and T. Conway. 2002. Gene expression profiling of the pH response in Escherichia coli. J. Bacteriol. 184:6551-6558.
48. Tucker, D. L., N. Tucker, Z. Ma, J. W. Foster, R. L. Miranda, P. S. Cohen, and T. Conway. 2003. Genes of the GadX-GadW regulon in Escherichia coli. J. Bacteriol. 185:3190-3201.
49. Vogel, H. J., and D. M. Bonner. 1956. Acetylornithinase of Escherichia coli: partial purification and some properties. J. Biol. Chem. 218:97-106.
50. Waterman, S. R., and P. L. C. Small. 1996. Characterization of the acid resistance phenotype and rpoS alleles of Shiga-like toxin-producing Escherichia coli. Infect. Immun. 64:2808-2811.
51. Waterman, S. R., and P. L. C. Small. 2003. Identification of the promoter regions and sigma-S-dependent regulation of the gadA and gadBC genes associated with glutamate-dependent acid resistance in Shigella flexneri. FEMS Microbiol. Lett. 225:155-160.
52. Yuan, J. T. C. 2003. Modified atmosphere packaging for shelf-life extension, p. 215-219. In J. S. Novak, G. M. Sapers, and V. K. Juneja (ed.), Microbial safety of minimally processed foods. CRC Press, Inc., New York, N.Y.(Arvind A. Bhagwat, Lynn C)