ArcA positively regulates the expression of virulence genes and contributes to virulence of porcine Shiga toxin-producing enterotoxigenic Escherichia coli

ABSTRACT

Enterotoxigenic Escherichia coli (ETEC) cause severe diarrhea in humans and animals. These bacteria encode two major classes of virulence factors, adhesins that promote colonization of the small intestine and enterotoxins that induce fluid secretion and thus diarrhea. Oxygen and redox sensors are regulators of virulence expression in multiple bacterial pathogens. In this study, we show that microaerobic conditions enhance the expression of virulence factors in a porcine Shiga toxin-producing ETEC (STEC/ETEC) strain. We then investigated the role of the aerobic respiration control regulator ArcA in the pathogenicity and virulence regulation in STEC/ETEC. In a mouse model, deletion of arcA caused less severe pathology, and the arcA mutant displayed lower levels of colonization and obvious weight gain of the mice compared to the wild type (WT). In a cell culture model, a lack of arcA reduced the adherence of STEC/ETEC to porcine intestinal epithelial cells. Furthermore, ArcA positively regulated the expression of several key virulence factors, including F18 fimbriae (fed), heat-labile (eltA) and heat-stable (estB) toxins, Shiga toxin 2e (stx2e), and hemolysin (hlyC), under microaerobic conditions and in vivo conditions. We then found that ArcA positively regulated the expression of eltA, estB, and hlyC by competing with a global repressor H-NS (histone-like nucleoid structuring). Mechanistically, we show that ArcA protein directly binds to the promoters of target genes, displaces H-NS silencing from the promoter and counteracts H-NS’s repression. Collectively, our data established a key role for ArcA in the pathogenicity and virulence expression of porcine STEC/ETEC.

IMPORTANCE

Enterotoxigenic Escherichia coli (ETEC) cause severe diarrhea in humans and animals, leading to death and huge economic loss worldwide. Thus, elucidation of ETEC’s pathogenic mechanisms will provide powerful data for the discovery of drugs serving as prevention or therapeutics against ETEC-caused diarrheal diseases. Here, we report that ArcA plays an essential role in the pathogenicity and virulence regulation in ETEC by positively regulating the expression of several key virulence factors including F18 fimbriae, heat-labile and heat-stable toxins, Shiga toxin 2e, and hemolysin, under microaerobic conditions and in vivo. Moreover, we found that positive regulation of several virulence genes by ArcA requires a global repressor H-NS (histone-like nucleoid structuring), implying that ArcA may exert positive effects by antagonizing H-NS. Collectively, our data established a key role for ArcA in the pathogenicity of porcine ETEC and ETEC strains isolated from human infections. Moreover, our work reveals another layer of regulation in relation to oxygen control of virulence factors in ETEC.

INTRODUCTION

Enterotoxigenic Escherichia coli (ETEC) is a prominent enteric pathogen responsible for causing tens of millions of cases of diarrheal illnesses annually. Its impact on young children, particularly those under the age of 5, is of significant concern due to their heightened vulnerability to ETEC infection, especially in regions where the pathogen is endemic. In 2015, ETEC was responsible for an estimated 100 million cases of diarrhea, a staggering figure that underscores the severity of the issue, tragically resulting in approximately 60,000 fatalities within this vulnerable age group (1, 2). ETEC is also a very common cause of diarrhea in agricultural production animals, including pigs, cattle, and sheep (3, 4). Diarrhea caused by ETEC in neonatal and post-weaning pigs is an economically significant disease worldwide due to mortality, weight loss, slow growth, and treatment cost (5). ETEC enter animals via the oral route usually in the form of contaminated food or water, and after passing through the stomach, these bacteria can attach to the intestinal epithelium using fimbrial or non-fimbrial adhesins. Adhesins frequently carried by animal/human ETEC include K88 (also called F4), F18, K99 (F5), 987P (F6), and F41. Once colonized, ETEC can multiply rapidly and secrete one or more classes of enterotoxins, such as heat-labile enterotoxin (LT) and heat-stable enterotoxin (ST). These toxins can induce the loss of water and electrolytes from epithelial cells into the gut lumen, and subsequently the onset of diarrhea (4, 6). Some animal/human ETEC strains carry Shiga toxin (Stx) type 2e, giving rise to hybrid Shiga toxin-producing enterotoxigenic E. coli (STEC/ETEC) (7, 8). Stx2 variant “e” encoded by the stx2eAB operon is the most frequent stx2 variant found in porcine fecal samples (9 – 11) and is highly associated with severe edema disease in pigs, which is also called edemigenic toxin (12). STEC/ETEC can induce both diarrhea and edema, leading to lethality in pigs (13, 14). Therefore, understanding its pathogenesis is of great significance and may aid in the development of therapeutics and antimicrobials.

Among all the adhesins, F18 is highly associated with strains causing porcine edema and post-weaning diarrhea (PWD) (10, 15, 16). F18 fimbriae is encoded by the fed (fimbriae associated with edema disease) gene locus comprised of six genes. The fedA gene encodes the major subunit of the fimbriae, fedB, the usher protein, fedC, the periplasmatic chaperone, fedE, a small linker protein between FedA and FedF, and fedF, the fimbrial adhesin (17, 18). FedF acts as an adhesin specific for porcine intestinal epithelial cells, and FedF is highly conserved in both F18ab and F18ac serological subtypes (19). In situations where the host is exposed to ETEC, and inhibitory milk glycans are absent after weaning, FedF has the capability to bind to a range of hexoses present in the glycosphingolipids of the intestinal cell membrane. This binding ability facilitates the initial attachment process (20).

Upon attachment, ETEC strains express one or both of the two classical toxins, LT and ST. Encoded by the eltAB operon, LT is a heterohexameric protein of ~84 kDa containing one A subunit (LTA) and five identical B subunits (LTB) (21). ST toxins are a group of low-molecular-weight enterotoxigenic peptides, usually containing 18 or 48 amino acids. STs are divided into two categories: the human and porcine (STp) ST based on their preferential hosts (22), or type 1 (ST1 or STa) and type 2 (ST2 or STb) ST based on their pathogenicity and solubility in methanol (23 – 25). STa and STb are encoded by estA and estB, respectively. Upon binding to their receptors, LT and ST trigger a cascade of cellular pathways, eventually leading to the transport of Cl⁻ and HCO₃ ⁻ to the intestinal lumen, as well as the loss of water (22, 26). ETEC strains may also possess hemolytic toxins, such as hemolysin (encoded by the hlyCABD operon) and cytolysin (encoded by clyA or hlyE) (27, 28). Although an ETEC strain containing only one adhesin and one enterotoxin is adequate to cause symptomatic diseases, hemolytic ETEC strains display increased virulence (29).

Previous studies demonstrate that the expression of some ETEC adhesins and enterotoxins are modulated in response to host environments, including pH, bile, and osmolarity (30). Two regulatory proteins CRP (cAMP-receptor protein) and H-NS (histone-like nucleoid structuring) play a key role in integrating extracellular signals such osmolarity and metabolism into the control of toxin production (31). A recent study shows that oxygen tension in the intestinal environment also influences the expression of genes encoding LT and the colonization factor antigen I (CFA/I) adhesin. In particular, under aerobic conditions, there was a substantial increase in the expression of LT and the CFA/I adhesion, in stark contrast to the anaerobic conditions. Intriguingly, there was no significant difference between the expression of these factors under aerobic and microaerobic conditions. However, it is worth noting that the expression of ST2 (heat-stable toxin 2) exhibited a significant increase under microaerobic conditions when compared to aerobic conditions. The global regulator FNR (fumarate and nitrate reduction) represses the expression of ETEC virulence factors (VFs) in the anaerobic lumen, and this repression can be relieved when ETEC nears the intestinal epithelium, where leaked oxygen from the epithelial cells inactivates FNR (32).

Similar to FNR, the Arc two-component signal transduction system, comprised of the kinase sensor ArcB and its cognate response regulator ArcA, enables E. coli adaptation to changing oxygen availability (33). ArcB kinase activity is modulated by the redox state of the ubiquinone and menaquinone pools (34). In response to low or no availability of oxygen, ArcB phosphorylates and activates ArcA, which then inhibits the expression of genes required for aerobic metabolism but stimulates genes implicated in the anaerobic respiratory and metabolic pathways (35, 36). Although the ArcA regulon has been extensively studied in E. coli K-12 strain (35 – 37), the role of ArcA for virulence and virulence regulation is less well studied in pathogenic E. coli. In this study, using a porcine hybrid STEC/ETEC strain (MD724-020) isolated from a piglet with acute diarrhea, we show that microaerobic conditions enhance the expression of VFs relative to anaerobic conditions and that ArcA positively regulates the expression of enterotoxins and hemolysin under microaerobic conditions, contributing substantially to the virulence of ETEC. Therefore, this work reveals another layer of regulation in relation to oxygen control of VFs in ETEC.

RESULTS

Microaerobic conditions enhance the expression of genes encoding enterotoxins and hemolysin

While the intestinal lumen is commonly thought to be anaerobic, the tissue surface surrounding the lumen can be microaerobic due to oxygen leaked from intestinal epithelial cells. Although accurate data about oxygen concentrations in the porcine intestine are lacking, noninvasive imaging of oxygen levels in the gastrointestinal tract of living mice indicated that the milieu in the mid-small intestine is microaerobic (1.4% of atmospheric pressure) (38). To assess whether the expression of genes encoding adhesin and toxin is different between microaerobic and anaerobic conditions, real-time quantitative PCR (RT-qPCR) assays were carried out with six selected genes in the porcine hybrid STEC/ETEC strain MD724-020 under both microaerobic (6%–12% O₂) and anaerobic (21% CO₂) conditions, which mimic in vivo conditions. As shown in Fig. 1, the expression of eltA, estB, stx2e, hlyA, and hlyE was significantly higher under microaerobic conditions than under anaerobic conditions, while the expression of the adhesin gene fedA was undetectable under both microaerobic and anaerobic conditions (data not shown). The largest fold changes were observed in stx2e and hlyC, with a >12 fold change and a >5 fold change, respectively. Given that the milieu environment in the mid-small intestine is likely microaerobic and the expression of several major virulence genes of porcine STEC/ETEC was higher in microaerobiosis than in anaerobiosis, microaerobic conditions were used for all expression assays and comparisons throughout this study.

Fig 1.

Microaerobic conditions enhanced the expression of enterotoxin and hemolysin genes. RT-qPCR analysis was performed to examine the expression of eltA, estB, stx2e, hlyC, and hlyE under microaerobic and anaerobic conditions. Relative expression levels were calculated using the tus gene as an internal control, and levels under anaerobic conditions were set as 1.0. Values represent the mean ± standard deviation of triplicate samples from three independent experiments. Significant differences were evaluated by Student’s t-test, and the asterisks indicate ** as P < 0.01 and * as P < 0.05.

Deletion of arcA significantly attenuates STEC/ETEC virulence both in vivo and in vitro

To address the role of ArcA in STEC/ETEC, the virulence of the wild-type MD724-020 ∆arcA mutant and ∆arcA complementation strains was compared using a 4-week-old mouse model. We first demonstrate that WT, ΔarcA mutant, and ∆arcA complementation strains grew at similar rates under aerobic, microaerobic, or anaerobic conditions (Fig. S1). Mice were pretreated with an antibiotic cocktail prior to infection. The WT, ∆arcA mutant, and ∆arcA complementation strains were inoculated into eight mice (n = 8 per group) with an inoculation dose of 1 × 10⁹ colony-forming units (CFU). We observed that three mice in the WT and two in the ∆arcA complementation strain-challenged group died within 15 days post-infection (dpi). The survivors displayed signs of distress and exhibited bloody stool. In stark contrast, in the group infected with the ∆arcA mutant, there were no recorded deaths or apparent symptoms at 15 dpi mark. Although the difference between the WT and ΔarcA mutant groups is not statistically significant (P = 0.0888, Fig. 2A), there is a trend that loss of arcA can increase the survival of mice. Additionally, the body weights of the surviving mice were monitored for 2 weeks. As shown in Fig. 2B, the WT and ∆arcA complementation strain-challenged mice showed significant weight losses from 2 to 15 dpi compared with the ∆arcA mutant-challenged mice (P < 0.01). The CFU analysis of fecal samples confirmed the absence of the STEC/ETEC strains before challenge. Fecal shedding of STEC/ETEC in each group was calculated by determining the CFUs of STEC/ETEC on blood agar, and no significant differences in shedding were observed between the ∆arcA and WT groups at 1 dpi. As shown in Fig. 2C, ∆arcA strain shedding was significantly lower than WT at 2 and 3 dpi (P < 0.01 and P < 0.05). Three days later, fecal shedding of most mice dropped to an undetectable level. Besides, histopathological examination detected pronounced changes in the WT and ∆arcA complementation strain-challenged mice but not in the mice challenged by the ∆arcA mutant. WT and ∆arcA complementation strain infection caused atrophy of microvilli and a large number of inflamed cells in the lamina propria of the ileum, but nearly no histological changes were observed in the mice infected with the ∆arcA mutant (Fig. 2D). These results demonstrate that the ∆arcA mutation significantly attenuates STEC/ETEC virulence in the murine model.

Fig 2.

ArcA contributes to the STEC/ETEC virulence. Groups of eight antibiotic-treated mice were challenged via oral gavage with 10⁹ CFU of the wild type, the ∆arcA mutant, or the ∆arcA complemented strains, and the survival (A), body weight change (B), fecal bacteria shedding (C), and intestine histopathology (D) were determined at different times post-infection. Data are presented as the mean ± standard deviation. Significant differences were evaluated by log rank test (A) and Student’s t-test (B and C), and the asterisks indicate * as P < 0.05, ** as P < 0.01, and *** as P < 0.001. ns, not significant. Hematoxylin and eosin staining images are representative of three replicates (magnification = 200). (E) Deletion of arcA significantly decreased ETEC adherence to porcine intestinal epithelial (IPEC-J2) cells. IPEC-J2 neonatal jejunal epithelial cells were infected with ETEC strains at a multiplicity of infection (MOI) of 10 for 1 h. After four washes with phosphate-buffered saline (PBS), bacteria associated with the cells were spread onto MacConkey agar plates and enumerated. Data are presented as the mean ± standard deviation of triplicate samples from three independent experiments. Significant differences were evaluated by one-way analysis of variance, and the asterisks indicate ** as P < 0.01.

To determine whether arcA affects the adherence of STEC/ETEC to porcine intestinal enterocytes isolated from jejunum, cultured porcine intestinal epithelial (IPEC-J2) cells were infected with the WT and ∆arcA mutant strains. As shown in Fig. 2E, deletion of arcA reduced STEC/ETEC adherence to IPEC-J2 cells by six-fold compared with that of the WT strains (P < 0.01). Complementation of the ∆arcA mutant by reintroduction of arcA greatly increased the levels of adherence. Therefore, these results suggest that arcA plays an important role in the adherence of STEC/ETEC to host cells.

ArcA regulates the expression of fedABCDEF encoding F18 pilus

The F18 pilus encoded by the fed gene cluster fedABCDEF mediates colonization of both ETEC and STEC in weaned pigs (39). We deleted the gene cluster fedABCDEF from the WT, and the adherence assay showed that a lack of the fedABCDEF operon dramatically reduced STEC/ETEC adherence to IPEC cells (Fig. 3A) (P < 0.01). To determine whether and how ArcA regulates the expression of fedABCDEF, we first used a reverse transcription-PCR assay to show that the gene cluster fedABCDEF forms one operon (Fig. 3B). The transcription start site (TSS) of the fedABCDEF operon was then mapped to the position 191 bp upstream of the fedA-coding region by using 5´-rapid amplification of cDNA ends (5´ RACE) PCR (Fig. S2), and putative −10 and −35 motifs of the promoter were identified as shown in Fig. 3C. Notably, when gene transcription levels were studied using chromosomal fedA-lacZ fusion and RT-qPCR under laboratory conditions in tryptic soy broth (TSB) and Luria-Bertani (LB) media, fedABCDEF expression was not detectable in the WT, ∆arcA mutant, or ∆arcA complementation strain. The hemagglutination assay results also revealed that both the wild-type strain and the ∆arcA complementation strains possess the capability to agglutinate chicken red blood cells, leading to their clumping together. In contrast, the ∆arcA mutant strain was found to be incapable of achieving this agglutination, as shown in Fig. 3D. During interaction with IPEC-J2 cells and infection in mice, the expression of fedABCDEF was significantly higher in the WT and ∆arcA complementation strains, than in the ∆arcA mutant (Fig. 3E and F), suggesting a positive role of ArcA in the expression of fedABCDEF during ETEC interaction with IPEC-J2 cells or during infection in vivo.The potential binding site of ArcA was identified with the Prodoric Virtual Footprint program (version 3.0) (40) at positions +15 to +24 downstream of the TSS of fedABCDEF (Fig. 3C). The ArcA-His₆ fusion protein was able to shift the DNA fragments containing the fedA promoter region but not the control fragment within the coding region (Fig. 3G). Altogether, these data suggest that ArcA likely regulates the expression of the fedABCDEF operon in a direct manner.

Fig 3.

ArcA positively regulated the expression of fedA. (A) Deletion of fedABCDEF significantly reduces ETEC adherence to porcine intestinal cells. (B) RT-PCR analysis indicates that the fedABCDEF genes are cotranscribed. RNA and cDNA samples were prepared as described above. Primers were designed to span open reading frames of fedA and fedBC, fedBC and fedD, fedD and fedE, and fedE and fedF, respectively. The black bars under the gene arrows denote the expected PCR amplicons, and the number in front of each bar corresponds to the lane in the agarose gel. The #1 fragment was used as a negative control reaction, as fedA and the gene encoding IS66 family transposase are not expected to cotranscribe. RNA that was not reverse transcribed served as a negative control template, while genomic DNA served as a positive control template. (C) Features of the fedA promoter region. Italic and bold, start codon; bold, transcription start site; shaded, −35 and −10 regions; and underlined, predicted ArcA binding sites. (D) Production of F18 was validated in WT, ∆arcA mutant, and ∆arcA complemented by chicken hemagglutination assay. (E) Expression levels of fedA during ETEC adhering to IPEC-J2 cells were assessed by RT-qPCR. (F) Expression of fedA during infection in the mouse intestine was compared in WT, ∆arcA, and ∆arcA complemented strains by RT-qPCR. Relative expression levels were calculated compared with the tus gene as an internal control, fold changes are relative to the WT. Data are presented as the mean ± standard deviation of triplicate samples from three independent experiments or from a pool of three mice. Significant differences were evaluated by one-way analysis of variance, and the asterisks indicate ** as P < 0.01 and * as P < 0.05. (G) Nonradioactive electrophoretic mobility shift assay (EMSA) showing phosphorylated ArcA-His₆ binding to the fedA promotor region. The PCR product of the fedA promoter region containing the ArcA binding site was used as a probe at 0.1 pmol per reaction mixture. A fedA-coding region fragment was used as a negative control. DNA fragments were stained with SYBR Gold.

Deletion of arcA significantly decreases the expression of heat-labile enterotoxin A in vitro and in vivo

The heat-labile enterotoxin A encoded by eltA is one of the well-known VFs for ETEC. To investigate whether and how ArcA regulates eltA expression, the TSS of eltA was mapped to the position 54 bp upstream of the eltA-coding region by 5´ RACE (Fig. S2), and the corresponding −10 and −35 motifs were identified at the proper distance to the TSS (Fig. 4A). The transcription of eltA was studied using chromosomal eltA-lacZ reporter gene fusion. Under microaerobic conditions, the expression of eltA was significantly reduced in the ∆arcA mutant in comparison to the WT (P < 0.01). Reintroduction of a plasmid carrying arcA restored eltA-lacZ expression at the transcriptional level (Fig. 4B). The expression of the EltA heat-labile toxin (EltA expression) in both the ΔarcA mutant and WT strains was assessed through Western blot analysis, employing a monoclonal antibody specific to EltA. As illustrated in Fig. 4C, the production of the EltA toxin exhibited a marked reduction in the ∆arcA mutant when compared to the WT strain, and this production was restored in the ∆arcA complemented strain (grayscale analysis was conducted using ImageJ for quantification). In addition, the expression of eltA during ETEC adherence to IPEC-J2 cells and during infection in mice was also compared in WT, ∆arcA, and ∆arcA complementation strains by RT-qPCR. As shown in Fig. 4D and E, the expression of eltA was significantly reduced in the ∆arcA mutant in comparison to that in the WT, and the expression levels increased in the ∆arcA complementation strain. These results indicate that ArcA regulates the expression of eltA under in vitro and in vivo conditions.

Fig 4.

ArcA positively regulated eltA expression. (A) Features of the eltA promoter region. Italic and bold, start codon; bold, transcription start site; shaded, −35 and −10 regions; and underlined, predicted ArcA binding sites. (B–E) A lack of arcA led to downregulation of eltA in media, during ETEC-host cell interaction and in vivo. (B) Expression levels of eltA were assessed by measuring β-galactosidase activities in eltA-lacZ transcriptional fusion strains grown microaerobically at 37°C in TSB medium. (C) The production of EltA was analyzed in WT, ∆arcA mutant, and ∆arcA complemented by Western blotting with GroEL as an internal control. (D) The expression of eltA during ETEC adherence to IPEC-J2 cells was assessed by RT-qPCR. (E) Expression of eltA during infection in the mouse intestine was compared in WT, ∆arcA, and ∆arcA complemented strains by RT-qPCR. Relative expression levels were calculated compared with the tus gene as an internal control, fold changes are relative to the WT. Data are presented as the mean ± standard deviation of triplicate samples from three independent experiments or from a pool of three mice. Significant differences were evaluated by one-way analysis of variance, and the asterisks indicate ** as P < 0.01 and * as P < 0.05. (F) An EMSA showing phosphorylated ArcA-His₆ binding to the eltA promoter region. This assay was performed according to Fig. 3G.

The potential binding sites of ArcA were identified by the Prodoric Virtual Footprint program (40). EMSAs showed that the ArcA protein shifted a DNA probe containing the potential ArcA binding sites but not the control DNA probe containing the coding region of eltA (Fig. 4F). Together, these data demonstrate that ArcA directly interacts with the eltA promoter and affects the expression of heat-labile enterotoxin gene A.

Deletion of arcA significantly decreases the expression of heat-stable toxin B in vitro and in vivo

The heat-stable toxin (ST) is another major enterotoxin produced by ETEC (24, 25), and the estB gene encodes STp (23). To investigate whether and how ArcA regulates estB expression, the TSS was identified by 5´ RACE PCR (Fig. 5A; Fig. S2), and the expression of estB was studied using chromosomal estB-lacZ reporter gene fusion. As shown in Fig. 5B, the expression of estB in the ΔarcA mutant was 2.6-fold lower than that in the WT under microaerobic conditions (P < 0.01). The production of STp heat-stable toxin (EstB expression) in the ΔarcA mutant and WT strains was further determined by enzyme-linked immunosorbent assay (ELISA) using a monoclonal antibody against STp. As shown in Fig. 5C, the production of STp toxin was significantly lower in the ∆arcA mutant compared with the WT, and the production was restored in the ∆arcA complemented strain. Furthermore, the expression of estB was significantly lower in the ∆arcA mutant compared with that in the WT (P < 0.01) during ETEC adhering to intestinal porcine enterocytes (Fig. 5D) and during infection in the mouse intestine (Fig. 5E). These results demonstrate that ArcA positively regulates the expression of heat-stable toxin B in vitro and in vivo.

Fig 5.

Deletion of arcA significantly decreased estB expression. (A) Features of the estB promoter region. Italic and bold, start codon; bold, transcription start site; shaded, −35 and −10 regions; and underlined, predicted ArcA binding sites. (B and C) A lack of arcA led to downregulation of eltA in TSB media. Transcription of estB-lacZ (B) and production of STb (C) were examined in WT, ∆arcA mutant, and ∆arcA complemented strains during growth in TSB media. Production of STb was analyzed by ELISA. (D) Expression levels of estB during ETEC adhering to IPEC-J2 cells were assessed by RT-qPCR. (E) Expression of estB during infection in the mouse intestine was compared in WT, ∆arcA, and ∆arcA complemented strains by RT-qPCR. Relative expression levels were calculated compared with the tus gene as an internal control, fold changes are relative to the WT. Data are presented as the mean ± standard deviation of triplicate samples from three independent experiments or from a pool of three mice. Significant differences were evaluated by one-way analysis of variance, and the asterisks indicate ** as P < 0.01 and * as P < 0.05. (F) An EMSA showing phosphorylated ArcA-His₆ binding to the estB promotor region.

The ArcA binding site in the promoter region of the estB gene was predicted (Fig. 5A), and the DNA fragments containing the potential binding sites were amplified by PCR. EMSAs showed that the ArcA protein shifted the DNA fragment containing the potential ArcA binding sites but not the negative control DNA fragments (Fig. 5F). These results suggest that ArcA could directly regulate the expression of the heat-stable toxin.

ArcA upregulates stx2e under in vitro and in vivo conditions

The ETEC strain used in this study possesses a stx2e gene encoding the edema toxin. The transcription start site of stx2e was determined by 5´ RACE PCR (Fig. S2), and the −35 and −10 motifs of the promoter were identified (Fig. 6A). The expression of stx2e was studied using chromosomal stx2e-lacZ reporter gene fusion. As shown in Fig. 6B, the expression of stx2e in the ΔarcA mutant was threefold lower than that in the WT under microaerobic conditions (P < 0.05). The regulation of Stx2e by ArcA was additionally validated through Western blot analysis. As illustrated in Fig. 6C, the Stx2e production in the ∆arcA mutant exhibited a notable and statistically significant decrease when compared to that in the WT (as depicted in Fig. 6C), with a P-value less than 0.01. The transcription of stx2e was also investigated in WT, ∆arcA, and ∆arcA complemented strains during ETEC adhering to intestinal porcine enterocytes (Fig. 6D) and during infection in the mouse intestine (Fig. 6E). Our results showed that stx2e was expressed at much higher levels in the WT and the complemented strains than in the ∆arcA mutant. These results indicate that ArcA positively affected the expression of stx2e. EMSAs further showed that the ArcA protein could bind to the promoter region of the stx2e gene (Fig. 6F), suggesting that ArcA may directly regulate the expression of stx2e.

Fig 6.

ArcA positively regulated stx2e expression. (A) Features of the stx2e promoter region. Italic and bold, start codon; bold, transcription start site; shaded, −35 and −10 regions; and underlined, predicted ArcA binding sites. (B–E) A lack of arcA led to downregulation of stx2e in media, during ETEC-host cell interaction and in vivo. (B and C) A lack of arcA led to downregulation of stx2e in TSB media. Transcription of stx2e-lacZ (B) and production of Stx2e (C) were examined in WT, ∆arcA mutant, and ∆arcA complemented strains during growth in TSB media. The production of Stx2e was analyzed by Western blotting with GroEL as an internal control. (D)The expression levels of stx2e during ETEC adherence to IPEC-J2 cells were assessed by RT-qPCR. (E) Expression of stx2e during infection in the mouse intestine was compared in WT, ∆arcA, and ∆arcA complemented strains by RT-qPCR. Relative expression levels were calculated compared with the tus gene as an internal control, fold changes are relative to the WT. Data are presented as the mean ± standard deviation of triplicate samples from three independent experiments or from a pool of three mice. Significant differences were evaluated by one-way analysis of variance, and the asterisks indicate ** as P < 0.01. (F) An EMSA showing phosphorylated ArcA-His₆ binding to the stx2e promotor region.

ArcA regulates the expression of hemolysin encoded by hlyA

Many ETEC strains encoding hlyCABD and/or hlyE are hemolytic (27, 41). For the STEC/ETEC strain used in this study, deletion of hlyA, but not hlyE, completely abolished the hemolytic activity, suggesting that hlyA, but not hlyE, is responsible for the hemolytic phenotype in the MD724-020 strain (Fig. 7A). To investigate whether ArcA regulates the expression of hlyCABD, the TSS of hlyCABD was obtained by 5´ RACE PCR (Fig. 7B; Fig. S2), and the transcription of the hlyCABD operon was studied using chromosomal hlyC-lacZ reporter gene fusion. As shown in Fig. 7C, the expression level of hlyC was significantly lower in the ∆arcA mutant compared with that in the WT (P < 0.01) when cultured in TSB under microaerobic conditions. The regulation of hlyCABD by ArcA was further confirmed by Western blot. As shown in Fig. 7D, the production of HlyA in the ∆arcA mutant was decreased by ~4-fold compared with that in the WT (grayscale as determined by ImageJ). Similarly, the expression of hlyC was significantly lower in the ∆arcA mutant compared with that in the WT during ETEC adhering to intestinal porcine enterocytes (Fig. 7E) and during infection in the mouse intestine (Fig. 7F) (P < 0.01). Furthermore, EMSAs were performed to test whether the ArcA regulator protein can bind to the promoter region of hlyCABD. As shown in Fig. 7G, the ArcA protein was able to shift DNA fragments containing the hlyCABD promoter region but not the negative control probe. Taken together, these results suggest that ArcA regulates the expression of the hlyCABD operon likely in a direct fashion, thus contributing to the hemolytic activity of ETEC.

Fig 7.

ArcA positively regulated hlyCABD expression. (A) Roles of hlyA and hlyE in hemolysis under microaerobic conditions, as revealed by a blood agar plate assay. (B) Features of the hlyC promoter region. Italic and bold, start codon; bold, transcription start site; shaded, −35 and −10 regions; and underlined, predicted ArcA binding sites. (C and D) A lack of arcA led to downregulation of hlyA in TSB media. Transcription of hlyA-lacZ (C) and production of HlyA (D) were examined in WT, ∆arcA mutant, and ∆arcA complemented and ∆hlyA mutant strains during growth in TSB media. The production of HlyA was analyzed by Western blotting with GroEL as an internal control. (E) Expression levels of hlyA during ETEC adhering to IPEC-J2 cells were assessed by RT-qPCR. (F) Expression of hlyA during infection in the mouse intestine was compared in WT, ∆arcA, and ∆arcA complemented strains by RT-qPCR. Relative expression levels were calculated compared with the tus gene as an internal control, fold changes are relative to the WT. Data are presented as the mean ± standard deviation of triplicate samples from three independent experiments or from a pool of three mice. Significant differences were evaluated by one-way analysis of variance, and the asterisks indicate ** as P < 0.01 and * as P < 0.05. (G) An EMSA showing phosphorylated ArcA-His₆ binding to the hlyCABD promotor region.

ArcA positively regulates eltA, estB, and hlyC by counteracting H-NS’s repression

Many activators upregulate gene expression by counteracting H-NS’s repression (42, 43). Previous reports have suggested that ArcA’s positive regulation of target genes may involve H-NS (44, 45). Using two single mutants, Δhns and ΔarcA, as well as a double mutant, ΔhnsΔarcA, we found that ArcA activates the expression of virulence genes (eltA, estB, and hlyC) but H-NS represses their expression, and that the transcription of these genes is comparable between Δhns and ΔhnsΔarcA (Fig. 8A through C), suggesting that positive regulation of these virulence genes by ArcA requires H-NS. Notably, regulation of stx2e by ArcA did not require H-NS, since the expression of stx2e was detectable in the WT (CT value 23.33 ± 0.85 of stx2e versus 22.39 ± 0.45 of tus), but not detectable in ΔarcA, Δhns, and ΔhnsΔarcA strains by RT-qPCR, suggesting a different regulatory mechanism. We then hypothesize that ArcA is able to displace H-NS from DNA in vitro, thereby relieving its repression. We first used EMSA to show that H-NS directly binds to the promoter regions of ArcA-regulated eltA (Fig. 8D). To examine competition between ArcA and H-NS, we set up reactions with both H-NS and ArcA added to the eltA promoter probe. These reactions contain a constant amount of H-NS but varying concentrations of ArcA. We observed that increasing concentrations of ArcA incrementally shifted the migration of the DNA-protein complex to resemble that of the ArcA-eltA complex (Fig. 8E). This result suggests that ArcA outcompete H-NS in binding to the promoter probe. To evaluate whether H-NS protein bound to the shifted DNA probe decreases and ArcA bound to the shifted DNA probe increases as ArcA concentration increases, EMSA reactions were transferred to nitrocellulose membrane such that the proteins bound to DNA probe could be monitored by immunoblotting. As shown in Fig. 8E, H-NS protein signal corresponding to the H-NS-eltA bound complex diminished, whereas ArcA signal intensified as ArcA concentration increased across reactions. Similar results were obtained for estB and hlyC (Fig. S3). Altogether, these results suggest that ArcA positively regulates eltA, estB, and hlyC by counteracting H-NS’s repression.

Fig 8.

Positive regulation of several virulence genes by ArcA requires H-NS. (A to C) Relative expression levels of eltA (A), estB (B), and hlyC (C) were assessed by RT-qPCR in WT, ∆arcA, ∆hns, and ∆arcA∆hns strains grown microaerobically at 37°C in TSB medium. Relative expression levels were calculated compared with the tus gene as an internal control, and levels in the WT were set as 1.0. Data are presented as the mean ± standard deviation of triplicate samples from three independent experiments. Significant differences were evaluated by Student’s t-test, and the asterisks indicate ** as P < 0.01 and * as P < 0.05. (D) An EMSA showing H-NS-FLAG binding to the eltA promotor region. (E) Competitive EMSA. Reactions containing 0.1 pmol eltA probe and 0.75 µM purified H-NS and either 0, 0.5, 1, 2, 3, 4, or 5 µM phosphorylated ArcA. Lanes labeled “–,” “H-NS,” “ArcA,” and H-NS (–DNA) indicate no protein, 0.75 µM H-NS, 2 µM ArcA, and 0.75 µM H-NS without DNA, respectively. The reactions were run on polyacrylamide gels and stained with 1 × SYBR Gold nucleic acid-staining solution (top panel) and then transferred to nitrocellulose membrane and probed for H-NS (middle panel) and ArcA (bottom panel) using anti-FLAG and for anti-His₆ antibodies, respectively.

DISCUSSION

The emergence of hybrid diarrheagenic Escherichia coli strains, which result from the fusion of genetic markers from different pathotypes, poses a global public health concern (46). This phenomenon is fueled by the presence of numerous virulence markers, often harbored on mobile genetic elements such as phages and plasmids. The mobility of these virulence genes via horizontal gene transfer fosters the evolution of hybrid pathotypes (47 – 51). STEC and ETEC are significant contributors to diarrhea in both humans and animals globally. Recent reports from diverse geographical regions, including Finland, Bangladesh, Sweden, and South Korea, have revealed the existence of hybrids resulting from the combination of STEC and ETEC strains, categorized as STEC/ETEC hybrids (7, 8, 52 – 54). Genome-wide phylogenetic analyses have shown a strong association between these hybrids and specific strains of ETEC and STEC, suggesting potential acquisition of Stx-phage and/or ETEC virulence genes during the evolution of STEC/ETEC hybrids (1, 10 – 12). The pivotal role of Stx-phages in facilitating gene transfer and driving the evolution of E. coli, including the development of STEC variants, cannot be overstated (55). Furthermore, plasmids carrying genes encoding ST and LT toxins have demonstrated their capacity to transfer among diverse E. coli strains (56). Notably, certain hybrid strains within this category have been epidemiologically linked to diarrheal diseases and hemolytic uremic syndrome in humans (7, 8, 53), underscoring the critical need for a comprehensive understanding of their virulence mechanisms. However, there remains a paucity of studies addressing the virulence and regulatory mechanisms of STEC/ETEC hybrid strains isolated from porcine sources. Therefore, this study aims to elucidate the virulence factors expressed by a hybrid STEC/ETEC strain and uncover the underlying pathogenic mechanisms employed by this unique pathogen.

Diarrhea caused by ETEC is very common in farm animals. ETEC strains produce two major types of VFs; adhesins that facilitate binding and colonization of host gut epithelium and enterotoxins that induce fluid secretion (4). To establish a successful infection, expression of VFs needs to be well coordinated. Here, we showed that several major VFs of STEC/ETEC are highly expressed under microaerobiosis, and the response regulator ArcA is required for full expression of the adhesin F18 and several other toxins, including ST and LT. F18 is a predominant adhesin of ETEC causing PWD and plays a crucial role in adherence to host cells (Fig. 3A); therefore, deletion of arcA impaired the adherence of ETEC to porcine intestinal epithelial cells (Fig. 2F). Among the affected toxins, ST and LT have been shown to affect ETEC virulence and/or colonization in mouse models (57, 58). Indeed, a lack of arcA severely reduced ETEC virulence in a mouse model. Further bioinformatic and experimental approaches demonstrate that ArcA antagonizes H-NS’s repression of target genes. Therefore, this study described the regulatory details of an important virulence regulator as well as its contributions to virulence in STEC/ETEC.

Interestingly, expression of the F18 fimbriae was not detectable under laboratory conditions. This is consistent with previous reports that not all strains express F18 in vitro. For example, most F18ab fimbriae of clinical isolates are produced at minimal levels on common laboratory media (59). Our results showed that expression of fed genes was detectable during in vivo infection or bacteria-host interaction. Similarly, a transcriptome study has revealed that several adhesins, such as type 1 fimbriae and a secreted autotransporter EatA, are activated following attachment of ETEC to intestinal epithelial cells (60). These data suggest that expression of ETEC VFs is tightly controlled and stimulated only when needed. However, the regulators involved in this process remain largely unknown. Here, we showed that the lack of arcA decreased the F18 expression to undetectable levels. Therefore, ArcA serves as a positive contributor in this host-adapted pattern of gene expression.

Besides host contact, oxygen is another important signal for human ETEC to regulate the expression of VFs, such as enterotoxins (32). This was reflected in our results that major ETEC VFs (ST, Stx2e, and hemolysin) are expressed at higher levels in microaerobiosis relative to anaerobiosis (Fig. 1). Microaerobic conditions also serve as inducing signals for virulence expression in some other enteric pathogens. In ETEC (61) and Shigella (62), type III secretion system (T3SS) can be induced by microaerobic conditions; likewise, in Salmonella, virulence genes on the virulence megaplasmid (pESI) are upregulated under microaerobiosis (63). Further, our results have suggested a positive role of ArcA in virulence regulation under microaerobiosis, and this role contrasted with the repressor FNR. Using a controlled human infection model for ETEC, Crofts et al. suggested that when inside the intestinal lumen where oxygen is very limited, expression of VFs are repressed by FNR. By contrast, as ETEC nears the epithelium, oxygen diffused from the epithelial cells can relieve FNR repression; and subsequently ETEC can colonize the tissue with the derepressed VFs (32). FNR and ArcA co-regulate multiple genes in E. coli K-12 (35), it seems highly likely that ArcA and FNR coordinate to optimal expression of virulence genes in ETEC. However, it is noteworthy that signals like salt and glucose concentration (30), and regulator proteins like CRP and H-NS (31) can modulate VF expression at other levels, thereby forming a regulatory network.

We also found that positions of the potential ArcA binding sites relative to the TSS are variable among the positively regulated genes (Fig. 3 to 7). These are in line with the findings made by Park et al., where they mapped the ArcA binding sites at a global scale and found that some binding sites are located downstream of TSS, while others occupy the −10 and/or −35 motifs. It has been suggested that activation by ArcA may not be through direct ArcA-RNA polymerase interaction, but rather through antirepression (35). Interestingly, our data indicate that H-NS, a global silencer on genes predominantly acquired through horizontal gene transfer (64), binds to the promoter regions of VFs and represses their expression; and that activation of eltA, hlyC, and estB by ArcA is dependent on H-NS. We further demonstrate that ArcA can displace H-NS from the promoter of target genes, thereby alleviating H-NS’s repression (Fig. 8). It seems not surprising that ArcA and H-NS compete for similar sites at a promoter, as both ArcA and H-NS binding sites on DNA are adenine-thymine (AT)-rich sequences (35, 65). Mechanistically, H-NS potentially causes bridge formation on DNA, thus blocking transcription initiation and/or elongation (66, 67); and eviction of H-NS from DNA by ArcA can lead to structural changes, e.g., opening of the complex, allowing transcription to start and continue. Similarly, Ler, a positive regulator of T3SS in enterohemorrhagic E. coli, competes for binding to the promoter region against H-NS and activates the expression of T3SS and virulence (43). Hence, our data reveal a previously unrecognized virulence counter-silencing mechanism in ETEC. But in the case of stx2e, ArcA positively regulates stx2e, independently of H-NS (Fig. S4), and this may be correlated to the distinctive binding site architecture (inverted repeats versus normally directed repeats). The molecular details underlying this regulatory mode warrant more future research.

In conclusion, the present work established that ArcA regulates multiple virulence genes under microaerobiosis and is required for full virulence in cell culture and mouse models. Therefore, ArcA can be added to the growing list of virulence regulators in STEC/ETEC. The discovery of drugs targeting ArcA may serve as therapeutic or prevention against STEC/ETEC-caused porcine diarrheal diseases.

MATERIALS AND METHODS

Bacterial strains, culture conditions, and plasmids

The wild-type strain STEC/ETEC MD724-020 (O141:H4) was isolated from the small intestine of a nursery-age pig that suffered from diarrhea and acute mortality in Maryland, United States, in May 2014 (68, 69). This particular strain was isolated through culturing on blood agar plates (5% sheep blood) and subsequently identified using matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Pathotyping of the strain was achieved through whole genome sequencing (WGS), revealing positive results for the presence of key genes including eltA, estB, stx2, stx2e, eastA, hlyCABD, hlyE, and the adhesin (F18) gene. The comprehensive WGS data have been deposited in the National Center for Biotechnology Information database under the Sequence Read Archive accession SRR7213616. The strains and plasmids used in this study are listed in Table S1. Aerobic growth was achieved by shaking in air at 160 rpm, while anaerobic and microaerobic culturing was conducted using AnaeroPack or MicaeroPack (Mitsubishi Gas Chemical Company, Japan) in a sealed jar. Growth of ETEC strains was measured in triplicate under aerobic, microaerobic, and anaerobic conditions for 12 h or 10 h, using a spectrophotometer. For genetic manipulations and β-galactosidase assays, all E. coli strains were routinely grown in TSB medium. Selective antibiotics and isopropyl-β-D-thiogalactopyranoside (IPTG) were added wherever necessary at the following concentrations: ampicillin, 50 µg/mL, kanamycin, 50 µg/mL, chloramphenicol, 25 µg/mL, nalidixic acid, 30 µg/mL, and IPTG, 0.1 mM.

Recombinant DNA techniques

PCR, DNA ligation, electroporation, and DNA gel electrophoresis were performed as described by Sambrook and Russell (70), unless otherwise indicated. Oligonucleotide primers were produced in the Iowa State University DNA facility (Iowa, USA) or purchased from BGI Tech Solutions (Guangzhou, China) and are listed in Table S2. All restriction and DNA-modifying enzymes were purchased from TaKaRa Biotechnology (TaKaRa, Dalian, China) and used according to the supplier’s recommendations. Recombinant plasmids, PCR products, and restriction fragments were purified using TaKaRa MiniBEST plasmid purification kits or agarose gel extraction kits (TaKaRa, Shiga, Japan) as recommended by the supplier. DNA sequencing was performed at the DNA facility of Shanghai Sunny Biotechnology Co., Ltd. Deletion mutants were constructed using the bacteriophage lambda red recombinase system described by Datsenko and Wanner (71). The mutants were confirmed by PCR and DNA sequencing.

Plasmid construction

For complementation, gene-coding sequences and their putative promoter regions were amplified using the primers listed in Table S2 from the WT strain and independently cloned into a low-copy plasmid pGEN-MCS (72) using BamHI and EcoRI sites. Chromosomal transcriptional lacZ fusion was constructed by homologous recombination of the suicidal plasmid pVIK112 carrying a fragment of the complete 5´-region of the target gene (73, 74). Briefly, PCR fragments of target genes were cloned into pVIK112 using EcoRI and XbaI sites. The resulting pVIK112 derivatives were introduced into STEC/ETEC MD724-020 ∆lacZ by conjugation. Conjugants were selected and confirmed by PCR. To construct the plasmid overproducing the ArcA-His₆ fusion protein, a 717 bp fragment containing the coding region of arcA was obtained by PCR from genomic DNA with primers carrying codons for 6 × His and subsequently cloned into the pET28a (+) vector (Novagen, Madison, WI, USA) using BamHI and HindIII sites. The resultant plasmid contained pET28a-ArcA-His₆ under the control of the T7 promoter. To construct the plasmid overproducing the H-NS-FLAG fusion protein, a 414 bp fragment containing the coding region of hns and 24 bp fragment coding for FLAG plus a stop codon were obtained by PCR and subsequently cloned into the pET21a (+) vector (Novagen). Constructs were verified by Sanger sequencing (Comate Corp., China).

Mapping of transcription start sites by 5´ RACE

Mapping of TSS was performed as previously described, but with minor modifications (75). Briefly, total RNA was isolated from wild-type MD724-020 after interaction with IPEC-J2 cells for 1 h. Genomic DNA was removed from the extracted RNA using the RNase-Free DNase set (Qiagen, California, CA, USA) and then reverse transcribed to cDNA using reagents supplied in the SMARTer RACE 5´/3´ Kit (TaKaRa, Shiga, Japan) following the manufacturer’s instructions. Random primers were used for first-strand cDNA synthesis. The obtained cDNA was then used for 5´ RACE amplification with Universal Primer A Mix and gene-specific primers (Table S2). The 5´ RACE products were cloned into the pRACE vector by in-fusion cloning. The fragments of several clones were sequenced for mapping of the primary transcription.

Purification of ArcA and H-NS proteins

Escherichia coli BL21 carrying pET28a-ArcA or pET21a-H-NS were grown in 200 mL of LB medium to an optical density at 600 nm (OD₆₀₀) = 0.6–0.8, and protein expression was induced by adding 0.1 mM Isopropyl β-D- Thiogalactopyranoside (IPTG) for 10 h at 28°C, respectively. The ArcA-His₆ fusion protein was purified as previously reported (76). The E. coli H-NS-FLAG protein was purified by Anti-DYKDDDK G1 Affinity Resin (GenScript, Piscataway, NJ, USA) according to the manufacturer’s protocol. Briefly, bacterial pellets after induction were resuspended in lysis buffer (50 mM Tris-HCl, pH 8.0, 150 mM NaCl, 1 mM phenylmethylsulfonyl fluoride (PMSF), 0.2 mg/mL DNase I, 10 mg/mL lysozyme) and incubated for 20 min, and then sonicated for 15 min. Supernatant was collected after centrifugation and loaded onto a 2 mL Anti-DYKDDDK G1 Affinity Resin column. Protein was eluted from the column using elution buffer (3 M NaCl). Fractions were analyzed by SDS-PAGE to test the presence and purity of H-NS-FLAG protein. Pooled fractions were concentrated using 10 kDa cutoff centrifugal filters (Sartorius) and dialyzed overnight against storage buffer (20 mM Tris-HCl, pH 8.0, 300 mM NaCl, 1 mM EDTA, 5% glycerol). Dialyzed protein was aliquoted and stored at −80°C.

EMSAs

To study the binding of ArcA to the DNA probe, EMSAs were performed as described previously. Briefly, DNA probes were amplified using specific primers and purified using a TaKaRa Mini-BEST gel extraction kit. EMSAs were performed by adding increasing amounts of purified and phosphorylated His₆-ArcA fusion protein (0 to 3.5 µM) to the DNA probe (0.1 pmol) in binding buffer (10 mM Tris-HCl, 7 mM MgCl₂, 5% glycerol, 40 µg/mL bovine serum albumin, pH 8.0) for 30 min at 37°C. The reaction mixtures were then subjected to electrophoresis on a 6% polyacrylamide gel in 0.5× Tris-borate-EDTA (TBE) buffer (44.5 mM Tris, 44.5 mM boric acid, 1 mM Ethylene Diamine Tetraacetic Acid (EDTA), pH 8.0) at 200 V for 45 min. The gel was stained in 0.5× TBE buffer containing 1× SYBR Gold nucleic acid-staining solution (Life Technologies, Grand Island, NY, USA) for 15 min, and then the image was recorded. To study the binding of H-NS to the DNA probe, binding reactions were performed in EMSA buffer (10 mM Tris-HCl, 50 mM KCl, 5 mM MgCl₂, 1 mM 1,4-dithio-DL-threitol (DTT), 1 mM EDTA, 5% glycerol, pH 8.0). Reactions were incubated at 37°C for 30 min. For competitive binding experiments, these reactions were then supplemented with the other protein (ArcA) in binding buffer and incubated at 37°C for an additional 15 min. The reaction mixtures were then subjected to electrophoresis on two separate 6% polyacrylamide gel at 200 V for 45 min using identical procedures. DNA was detected by staining the gels with 1× SYBR Gold nucleic acid-staining solution for 15 min at room temperature. To detect H-NS and ArcA proteins in competitive EMSAs, each stained gel was transferred to a nitrocellulose membrane. Proteins were then detected using anti-His-KPL (Sigma) and anti-FLAG-KPL (Sigma) antibodies.

β-galactosidase activity

Expression levels of the gene-lacZ reporter fusions were determined by measuring β-galactosidase activity of exponentially growing cultures at 37°C under microaerobic conditions in TSB medium. Overnight TSB cultures of E. coli containing the fusions of the gene of interest with lacZ were washed with phosphate-buffered saline, diluted 1:100 in TSB medium and grown at 37°C to log phase or stationary phase. These cultures were diluted 1:1 in Z buffer and assayed for β-galactosidase activity using ortho-nitrophenyl-β-galactoside as the substrate as described previously (77). The values of β-galactosidase activity were measured at least in triplicate for each experiment.

Western blot

Bacteria were cultured in TSB under microaerobic conditions at 37°C as described above and pelleted by centrifugation at 12,000 × g for 2 min. For EltA and Stx2e, cell pellets were resuspended in phosphate-buffered saline (PBS) and subjected to sonication and then centrifuged at 5,000 × g for 10 min to collect the supernatant. For HlyA, cell pellets were resuspended in double-distilled water and mixed with loading buffer. After being lysed by boiling for 10 min, cell lysates were centrifuged at 12,000 × g for 2 min at 4°C. Then, the supernatant was subjected to SDS-PAGE. Proteins were transferred to a polyvinylidene fluoride membrane (Millipore), and the membrane was blocked with 5% skim milk at 37°C for 2 h. Blots were probed with a polyclonal antibody against EltA (Abcam, ab243102), Stx2e (Abmart, China), HlyA (Abclonal, China), or against GroEL (Abcam, ab90522) and an anti-rabbit IgG KPL DyLight 800-labeled antibody (SeraCare, 5230-0346). Membranes were viewed by using an Odyssey CLx imager (Odyssey LICOR, USA).

STb-ELISA

The level of STb in cell-free culture supernatants was determined by STb-ELISA performed essentially as described previously with minor modifications (78). Briefly, crude culture supernatants (50 µL) for each strain from the brain heart infusion broth supplemented with 2% casamino acids (BHI-CA) media were diluted 1:2 in 0.1 M sodium carbonate buffer (pH 9.6) and used to coat the wells of 96-well flat bottom microtiter plates (Corning Incorporated, Kennebunk, ME). Each strain was coated onto six wells, and wells that received only carbonate buffer served as background control wells. The plates were coated overnight at 4°C, after which, the overnight coating solution was removed, and the plates were washed three times with 0.05% PBS and tapped dry. The remaining unbound reaction sites were blocked by the addition of 200 µL 5% skim milk to all wells and incubated for 1 h at 37°C. After removing the blocking solution and washing plates three times with PBS, 100 µL of mouse anti-STb toxin antibody (Magic Anti-E. coli Heat Stable Enterotoxin Monoclonal antibody, Clone N230630) was added to all wells, and plates were incubated for 1 h at 37°C. The plates were washed three times with PBS and tapped dry, and 200 µL of goat anti-mouse IgG alkaline phosphatase conjugate (1:2,000 in PBS; Abcam) was added to all wells and incubated for 1 h at 37°C. The plates were again washed three times with PBS and tapped dry. Then, 50 µL of 3,3´,5,5´-tetramethylbenzidine (Sigma-Aldrich, Shanghai, China) was added to all wells, and the plates were incubated in the dark at 37°C for 15 min. The reaction was stopped by the addition of 50 µL of 2 M H₂SO₄. The optical density of each reaction mix was measured at 450 nm with a plate reader (BioTek, Winooski, VT, USA).

Hemolysis assay

To assess hemolysis on blood agar (79), we streaked the WT, ∆hlyA, and ∆hlyE strains onto blood agar plates containing 10% defibrinated sheep blood. These plates were then incubated at 37°C for 18–24 h under microaerobic conditions. The extent of hemolysis was determined by the size and translucence of the hemolytic halo—larger or more translucent halos indicated stronger hemolytic activity, while smaller or less translucent halos indicated weaker activity.

Hemagglutination assay

To evaluate the bacterial cell surface expression of F18 fimbriae, we employed a hemagglutination assay using the WT, ∆arcA mutant, and ∆arcA complemented strains. This assay utilized fresh bacterial suspensions and chicken red blood cells, with a negative control consisting of a saline solution (0.9% NaCl). The hemagglutination tests were conducted in V-bottom microtiter plates. We prepared a series of twofold dilutions of the bacterial isolates in separate microtiter wells using saline solution. To each well containing the bacterial dilutions, we added equal volumes of washed red blood cells. After gently mixing, the microtiter plate was incubated at 37°C for 1 h.

Adherence assays

Cell adherence assays were performed as previously described (80). The neonatal jejunal epithelial cell line IPEC-J2 was cultured in Dulbecco’s Minimal Essential Medium basic (Gibco, Grand Island, NY, USA) containing 10% fetal bovine serum (Gibco, Grand Island, NY, USA) at 37°C in 5% CO₂. Cells were plated in sterile 24-well plates at 6 × 10⁴ cells per well 24 h before the experiment. STEC/ETEC MD724-020 wild-type, ∆arcA mutant, and complemented strains were cultured statically in TSB medium until the OD₆₀₀ was 0.6. Before infection, two wells of cultured cells were trypsinized, and the cells were counted to estimate the cell number per well. Cells were washed twice with 1 mL PBS and were then exposed to bacteria at a multiplicity of infection of 10 and washed once with PBS. The 24-well plates were centrifuged (500 × g for 5 min) and incubated for 1 h. For the adherence assays, bacterium-exposed cells were washed four times with 1 mL of sterile PBS and then lysed with 1 mL 0.1% Triton X-100 for 5 min at room temperature. Serial dilutions of cell suspensions were spread onto MacConkey agar plates (Becton Dickinson & Co, Franklin Lakes, NJ), and CFU counts were obtained after overnight growth at 37°C. The input dilution of bacteria was also plated to determine the CFU count for each inoculum.

Mouse infection by ETEC

Healthy 4-week-old BALB/c mice were used as the ETEC infection model (31, 81). All animals were allowed free access to sterile water and animal feed and were handled according to the guidelines of the Laboratory Animal Monitoring Committee of Heilongjiang Province. Mice were randomly divided into three groups according to the experiment with four female and four male mice in each group. In this study, the three groups were challenged with WT, ∆arcA mutant, or ∆arcA complementation strains. Following a 2-day acclimation period upon arrival, mice were given gentamicin (35 mg/L), vancomycin (45 mg/L), metronidazole (215 mg/L), and colistin (850 U/mL) in drinking water to disrupt resident microbiota as previously published (82). After 3 days on antibiotics in water, mice were given untreated water for 1 day. Each mouse was weighed and recorded 0 days before infection and then given a single oral challenge by gavage of 1 × 10⁹ CFU ETEC in 100 µL of a 1.2% carbonate solution. Feces were collected from infected mice, and the fecal bacterial load of each mouse was detected by blood agar plate counting. After the challenge, mice were monitored and weighed daily for a total of 2 weeks. When the animal test was completed, alive mice were humanely euthanized.

Histology

Mice were grouped and challenged with ETEC strains as described above. At 24 h post-infection, mice were sacrificed to collect the jejunum and ileum. Tissues were fixed with 4% paraformaldehyde and stained with hematoxylin and eosin.

RT-PCR and quantitative real-time RT-PCR

RT-PCR was used for the co-transcription test. RNA of the wild-type MD724-020 during interaction with IPEC-J2 cells was extracted using an RNeasy Mini Kit (Qiagen) with 1-h in-tube DNase digestion (Qiagen) to remove possible DNA contamination according to the manufacturer’s instructions. One microgram of total RNA was reverse transcribed in triplicate using random hexamers and ImProm-II reverse transcriptase (Promega, Madison, WI, USA). For RT-PCR, primer pairs were designed to span adjacent genes. cDNA was then used as the template for subsequent PCRs, and RNA that was not reverse transcribed was used as a negative control.

RT-qPCR was used to validate the expression levels of selected genes. For the in vivo mouse infection test, total RNA was extracted from the intestinal contents of mice infected with WT, ∆arcA mutant, or ∆arcA complementation strains using the E.Z.N.A Stool RNA Kit (Omega, Norcross, GA, USA), and non-infected mice were included as a control. For the in vitro cell infection test, IPEC-J2 cells were previously infected with WT, ∆arcA mutant, or ∆arcA complementation strains for 1 h. Then, the medium was removed, and the cells were washed gently three times with PBS and resuspended in 0.1% PBST (phosphate-buffered saline with 0.05% Tween 20). The mixture of cells and bacteria was centrifuged for 5 min at ×1,000 g, the supernatant was collected and centrifuged for 2 min at ×12,000 g, and then the collected bacteria were used for RNA extraction using the SV Total RNA Isolation System (Promega, Madison, WI, USA). Total RNA from in vivo and in vitro experiments was treated with gDNA Eraser and reverse transcribed with random primers and reverse transcriptase using the PrimerScript RT reagent Kit with gDNA Eraser (TaKaRa). The primer pairs used are listed in Table S1 in the supplemental material, and RT-qPCR was performed as previously described (76, 83). Melting-curve analyses were performed after each reaction to ensure amplification specificity. The Ct value was set as 38 when gene expression was below detectable level. Differences (n-fold) in transcripts were calculated using the relative comparison method, and the amplification efficacies of each primer set were verified as described by Schmittgen et al. (84). RNA levels were normalized using the housekeeping gene tus for the replication terminator protein as a control.

Statistical analysis

Figures were plotted with GraphPad Prism 8 software. The significance of differences between groups was evaluated using SPASS statistics version 20 (SPASS Software). One-way analysis of variance was used when the sample size was greater than 2, and Student’s t-test was applied for two-group analysis. Differences were considered significant at a P-value of 0.05.

ETHICS APPROVAL

Handling and Care of mice were performed according to the Beijing Administration Guidelines for the Use of Laboratory Animals. The entire protocol with respect to animal experiments was approved by the Review Board of Harbin Veterinary Research Institute and by the Animal Care and Use Committee of Heilongjiang Province (SYXK(H)2006–032). All efforts were made to minimize suffering of animals.

SUPPLEMENTAL MATERIAL

The following material is available online at https://doi.org/10.1128/spectrum.01525-23.

Supplemental material. spectrum.01525-23-s0001.pdf.

Fig. S1 to S4; Tables S1 and S2.

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