Research Article
19 December 2022

Omptin Proteases of Enterobacterales Show Conserved Regulation by the PhoPQ Two-Component System but Exhibit Divergent Protection from Antimicrobial Host Peptides and Complement


Bacteria that colonize eukaryotic surfaces interact with numerous antimicrobial host-produced molecules, including host defense peptides, complement, and antibodies. Bacteria have evolved numerous strategies to both detect and resist these molecules, and in the Enterobacterales order of bacteria these include alterations of the cell surface lipopolysaccharide structure and/or charge and the production of proteases that can degrade these antimicrobial molecules. Here, we show that omptin family proteases from Escherichia coli and Citrobacter rodentium are regulated by the PhoPQ system. Omptin protease activity is induced by growth in low Mg2+, and deletion of PhoP dramatically reduces omptin protease activity, transcriptional regulation, and protein levels. We identify conserved PhoP-binding sites in the promoters of the E. coli omptin genes ompT, ompP, and arlC as well as in croP of Citrobacter rodentium and show that mutation of the putative PhoP-binding site in the ompT promoter abrogates PhoP-dependent expression. Finally, we show that although regulation by PhoPQ is conserved, each of the omptin proteins has differential activity toward host defense peptides, complement components, and resistance to human serum, suggesting that each omptin confers unique survival advantages against specific host antimicrobial factors.


Enterobacterales are an order of Gram-negative bacteria that commonly act as commensals in the mammalian gastrointestinal tract but also include numerous important pathogens, including Klebsiella pneumoniae, Salmonella enterica, and Escherichia coli. Escherichia coli is a common cause of acute gastroenteritis as well as extraintestinal infections, including urinary tract infections and sepsis (1, 2). Bacteria that colonize the human gastrointestinal tract or other mucosal surfaces encounter a number of environmental stressors, and they have evolved sophisticated mechanisms for both detecting these stressors and for mounting an appropriate defensive response. These bacterial defense systems are critical for the lifestyle of the microbe, and understanding how these systems are regulated and how they function is an important research goal.
In order to protect themselves from infection, mammalian hosts have developed a sophisticated system of both innate and adaptive immune defenses. The innate immune system is the first line of defense and includes both cellular components like neutrophils and macrophages as well as soluble molecules with direct antimicrobial and/or innate pattern recognition activity. Soluble components of the innate immune response include cationic antimicrobial peptides, secreted pattern recognition receptors, and components of the complement signaling cascade. In the complement pathway, serum components recognize conserved structures on the surfaces of microbial cells, leading to the formation of a membrane attack complex and bacterial lysis (3). The pathway also serves to recruit cellular innate immune components, thereby leading to enhanced infection control. In spite of this, numerous bacteria have evolved mechanisms to resist the activity of complement in order to enhance virulence (4). Mechanisms of complement resistance include preventing opsonization, promoting binding of soluble complement regulatory factors, or proteolytic degradation of components of the complement cascade (5).
Cationic antimicrobial peptides are a structurally diverse group of molecules that contribute to both constitutive and inducible host defense (6). These peptides bind to and penetrate bacterial membranes, leading to localized aggregation, reorganization, and disruption (7). This membrane activity is thought to be the main mechanism by which the majority of cationic antimicrobial peptides function. An important antimicrobial host defense peptide (HDP) is LL-37. This peptide is a cathelin family peptide and is produced by neutrophils and macrophages as well as by mucosal epithelial cells throughout the body (8). During inflammation, it is upregulated in epithelial cells and is also secreted at higher levels by infiltrating cells of the innate immune system (9). LL-37 has broad immunomodulatory effects as well as direct antibacterial effects, principally by interactions with the bacterial cell envelope leading to loss of cellular integrity and cell death (1012). Other mammals express different cathelicidin proteins, including mCRAMP in mice, protegrin in pigs, and bactenecin and indolicidin in cows (13). Together, complement family proteins and cationic antimicrobial peptides protect mucosal and endothelial surfaces from potential microbial attack.
Bacteria are not willing participants in this host-mediated defense system, however. In the Enterobacterales, many species have evolved regulatory systems that respond directly to environmental cues, including changes in external osmolarity or pH, exposure to host- or bacterial-derived signaling molecules, and the presence of host defense peptides themselves (1416). In E. coli, the best-characterized mechanism for resistance to host defense peptides involves two separate two-component signaling pathways, the PhoPQ and PmrAB systems. The PmrAB system responds directly to high concentrations of Fe3+ and regulates a number of genes that modify the bacterial surface charge by adding either 4-aminoarabinose or phosphoethanolamine to anionic phosphate groups in the lipid A and/or core region of lipopolysaccharide (LPS) (15). The PhoPQ system responds to limiting concentrations of divalent cations (17) or to antimicrobial peptides themselves (18). The PhoPQ system can modify the acylation status of lipid A via the regulation of the PagP palmitoyltransferase (19). Importantly, PhoPQ transcriptionally regulates the pmrD gene and PmrD posttranscriptionally activates PmrA, leading to expression of PmrA-regulated genes. Thus, the net effect of activating the PhoPQ signaling system is that the bacteria become resistant to cationic host defense peptides due to reduced peptide binding and altered membrane hydrophobicity.
In addition to reducing surface peptide binding, bacteria may also express outer membrane-associated proteases that can degrade proteins that are in the bacteria or on the bacterial surface (20). These omptin family proteins, named for the prototypical OmpT protein, are 10-stranded β-barrels in which the protease catalytic site is found in the surface-associated loops (21). The catalytic site for omptins includes a strictly conserved His-Asp dyad that activate a nucleophilic water molecule and an Asp-Asp dyad that is believed to stabilize the intermediate state (22). Omptin proteases exhibit a strong preference for cleavage at dibasic sites, although some exceptions to this exist (23). This preference appears to be due to a fairly anionic pocket at the base of the loop region and adjacent to the catalytic site (21). Numerous OmpT homologs are found throughout the Enterobacterales, including PgtE in Salmonella, CroP in Citrobacter, OmpP and ArlC (24) in E. coli, SopA in Shigella, and Pla of Yersinia pestis (25). These proteins can cleave some host defense peptides, and they contribute to resistance to this class of molecules in numerous bacterial species (26). Here, we demonstrate that all known omptins in E. coli are directly regulated by the PhoPQ system. We further show that each omptin orthologue provides differential protection from antimicrobial host defense peptides and from complement-mediated killing. The ArlC omptin protease is the most effective at protecting bacteria from LL-37-, CRAMP-, and whole-human-serum-mediated killing. These results confirm predictions about the role of PhoP in omptin regulation and demonstrate that ArlC carriage is an important host resistance factor for E. coli.


Expression and activity of ompT are regulated in a limiting Mg2+- and PhoP-dependent manner.

We monitored cleavage of an LL-37-derived Förster resonance energy transfer (FRET) substrate by the E. coli K-12 strain BW25113 and the ΔphoP and ΔpmrA derivatives thereof grown under conditions that would differentially activate PhoP or PmrA (low [10 μM] Mg2+ and low [1 μM] Fe3+), PmrA only (high [10 mM] Mg2+ and high [100 μM] Fe3+), or neither system (high [10 μM] Mg2+ and low [1 μM] Fe3+). As seen in Fig. 1A, growth of BW25113 under low-Mg2+ conditions resulted in an ~8-fold induction of protease activity relative to growth in N-minimal medium containing high Mg2+. In contrast, growth in the presence of high Fe3+ and high Mg2+ did not induce this activity. We next created plasmid-based translational ompT-green fluorescent protein (GFP) fusions and assessed fluorescent activity after growth under low- and high-Mg2+ conditions. As observed in Fig. 1B, low Mg2+ resulted in significant increases in GFP signal. Production of OmpT protein, as assessed by Western blotting, also followed the same response to different Mg2+ concentrations (Fig. 1C and E). Low Mg2+ but not high Fe3+ resulted in ~6-fold upregulation of the OmpT protein relative to the control DnaK control. This shows that PhoPQ-inducing conditions increased OmpT gene and protein expression as well as protease activity. Growth under PmrAB-inducing conditions, however, had little effect.
FIG 1 OmpT activity and expression are upregulated during growth under Mg2+ conditions in a PhoP-dependent manner. (A) Förster resonance energy transfer (FRET)-based activity of whole-cell, mid-logarithmic-phase culture of BW25113 grown in N-minimal medium with low Mg2+ and low Fe3+ (10 μM Mg2+ and 1 μM Fe3+), high Mg2+ and low Fe3+ (10 mM Mg2+ and 1 μM Fe3+), or high Mg2+ and high Fe3+ (10 mM Mg2+ and 100 μM Fe3+) using an LL-37-based probe as described in Materials and Methods. (B) A plasmid-based ompT-GFP transcriptional reporter fusion was transformed into BW25113, BW25113 ΔpmrA, and BW25113 ΔphoP, and the fluorescence was examined during growth in N-minimal medium containing low Mg2+ (10 μM) or high Mg2+ (10 mM). (C) Western blotting for OmpT proteins prepared from mid-logarithmic-phase-grown cultures of BW25113, BW25113 ΔpmrA, and BW25113 ΔphoP in N-minimal medium containing 10 μM Mg2+; (D) Western blotting for OmpT proteins prepared from mid-logarithmic-phase-grown cultures under the conditions indicated; (E) densitometric quantitation of Western blots for OmpT protein prepared from mid-logarithmic-phase-grown cultures of BW25113, BW25113 ΔpmrA, and BW25113 ΔphoP in N-minimal medium containing 10 μM Mg2+; (F) densitometric quantitation of Western blots for OmpT proteins prepared from mid-logarithmic-phase-grown cultures under the conditions indicated. The results shown are the mean and standard error of the values from each treatment. Statistics indicate P values from two-way analysis of variance (ANOVA) with Dunnett’s multiple-comparison testing. ns, no statistical difference; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.
To further probe the role of PhoP in OmpT regulation, we compared BW25113 to isogenic ΔpmrA or ΔphoP mutants. As seen in Fig. 1A, we observed a dramatic reduction in FRET activity in low-Mg2+-induced activity in the ΔphoP mutant. The ΔphoP mutant similarly exhibited very little ompT gene expression (Fig. 1B). Consistent with this, loss of functional PhoP also resulted in significant loss of OmpT protein as assessed by Western blotting (Fig. 1D and F). Both OmpT activity and expression were observed in the pmrA mutant when PhoPQ was induced with low Mg2+. Taken together, these results suggest that the PhoPQ system plays a role in OmpT regulation, and this is not dependent on the downstream regulatory system PmrAB.

Omptin activity in diverse Enterobacterales strains is also PhoP dependent.

In E. coli, three omptin proteins, OmpT, OmpP, and ArlC, have been described to date, and these have been linked to degradation of cationic molecules in the bacterial environment (24, 2729). In addition, promoter diversity in the ompT gene has also been associated with elevated omptin production in enterohemorrhagic E. coli (EHEC) O157:H7 (28). Salmonella enterica and Citrobacter rodentium are other Enterobacterales that encode an OmpT orthologue (PgtE and CroP, respectively). In order to test the PhoP dependence of omptin activity in these strains, we obtained isogenic PhoP or PhoPQ mutants of each strain and subjected the strains to FRET assay. As seen in Fig. 2, deletion of the phoP gene from three additional E. coli strains—adherent-invasive E. coli (AIEC) strain LF82 (24, 30), AIEC strain NRG857c (24, 31), and EHEC strain 86-24 (32)—also abrogated omptin activity in these strains. We also obtained C. rodentium DBS100 and an isogenic phoPQ mutant (33). Consistent with what was observed in all of the E. coli strains, the C. rodentium ΔphoPQ mutant also showed abrogation of omptin protease activity.
FIG 2 Multiple strains of Escherichia coli and Citrobacter rodentium exhibit PhoP-dependent omptin activity. Förster resonance energy transfer (FRET)-based activity of whole-cell, mid-logarithmic-phase cultures of either WT (black boxes) or ΔphoP mutant (white boxes) strains of each of the bacterial strains indicated. The results shown are the mean values from three independent biological replicates of the samples as indicated.

PhoP-dependent regulation of OmpT orthologues OmpP, ArlC, and CroP.

Our observation of different omptin activities in strains/species containing diverse omptin orthologues in multiple species of Enterobacteriales led us to test whether the regulation observed for ompT-GFP was also conserved in these other omptin orthologues. We constructed a series of GFP reporter fusions, including ompP-GFP, arlC-GFP, and croP-GFP. These plasmid-encoded fusions were then transformed to appropriate host strains, including E. coli BW25113 or BW25113 ΔphoP and C. rodentium DBS100 or DBS100 ΔphoPQ. For all omptins tested, expression was induced in both the wild-type (WT) and ΔpmrA strains under low-Mg2+ conditions (Fig. 3). Omptin expression was dramatically reduced in the phoP mutant strains. Taken together, these results are consistent with omptin expression that is controlled in a PhoP-dependent manner.
FIG 3 Diverse omptin orthologues are transcriptionally regulated by limiting Mg2+ in a PhoP-dependent manner. A plasmid-based omptin-GFP transcriptional reporter fusion was transformed into BW25113, BW25113 ΔpmrA, and BW25113 ΔphoP to measure (A) ompP, (B) arlC, or (C) croP expression. The fluorescence was examined during growth in N-minimal medium containing low Mg2+ and low Fe3+ (10 μM Mg2+ and 1 μM Fe3+), high Mg2+ and low Fe3+ (10 mM Mg2+ and 1 μM Fe3+), or high Mg2+ and high Fe3+ (10 mM Mg2+ and 100 μM Fe3+) as indicated in the figure. The results shown are the mean and standard error of the values represented in each treatment. Statistics indicate P values from two-way ANOVA with Dunnett’s multiple-comparison testing. ns, no statistical difference; ***, P < 0.001; ****, P < 0.0001.

PhoP-dependent regulation of OmpT requires a conserved PhoP-DNA binding site.

Genes that are directly PhoP regulated contain a conserved PhoP-binding motif, termed the PhoP box (34). We identifed putative PhoP boxes in the promoters of ompT, ompP, and arlC of E. coli as well as in croP of C. rodentium (Fig. 4A). The sites were located upstream of putative −35 and −10 sites as identified by the BPROM algorithm (35). We mutated the putative PhoP-binding site in our ompT-GFP reporter fusion from CTAAACAAAATATAAACAG to CTAAACAAAATATGGGCAG (underlined portions represent the PhoP box as shown in Fig. 4A. The bold portion represents the specific mutation introduced into the ompT HS promoter [as shown in Fig. 4B]). As seen in Fig. 4C, this mutation resulted in a significant loss of low-Mg2+ and PhoP-dependent induction of ompT-GFP activity, consistent with this PhoP-binding site being critical for low-Mg2+-induced ompT transcriptional activity.
FIG 4 Identification of a PhoP-binding site required for low-Mg2+-regulated ompT-GFP fusion activity. (A) Identification of conserved putative PhoP-binding sites (yellow highlight) in omptin family proteins of Escherichia coli (encoded by ompT, ompP, and arlC) and Citrobacter rodentium (encoded by croP). The identities of putative −10 and −35 sites, as identified by the BPROM promoter finding algorithm (cyan highlight), and the transcriptional (underlined) and translational (green highlight) start sites of ompT are also shown. (B) Sequence of the engineered AAA→GGG half-site (HS) mutation (red highlight) in the ompT-GFP promoter; (C) fluorescence of ompT-GFP or ompT-HS mutant-GFP fusions in N-minimal medium containing low Mg2+ (10 μM) or high Mg2+ (10 mM) as indicated in the figure; (D) PhoP-HA pulldown assay. Biotin-tagged DNA-probe and streptavidin bead complexes were treated with lysates of cultures grown to mid-log phase in low Mg2+ (10 μM). PhoP was detected using anti-HA antibody following SDS-PAGE and membrane transfer. Quantification of five replicates, each normalized to the no-probe control, is shown in panel E. The results shown are the mean and standard error of the values represented in each treatment. Statistics indicate P values from one-way ANOVA with Tukey’s multiple-comparison testing. ns, no statistical difference; *, P < 0.05; **, P < 0.01; ****, P < 0.0001.
We next probed PhoP-DNA interaction using a pulldown assay with biotinylated DNA probes that could be purified using streptavidin-agarose beads. When exposed to cell lysate containing hemagglutinin (HA)-tagged PhoP, this DNA probe-streptavidin bead complex would also purify any bound PhoP along with it. This interaction was then visualized with anti-HA antibody, which binds to the HA-tagged PhoP protein in the cell lysate. We observed that the WT ompT promoter bound significantly more PhoP-HA than the ompT promoter where one of the half-sites was mutated (Fig. 4D and E). The promoter for the pmrD gene, which is a known target of PhoP, also similarly bound PhoP-HA. A negative control, consisting of a reaction mixture containing the dnaK promoter, showed background levels of PhoP-HA, similar to a reaction mixture where there was no biotinylated probe added. This demonstrates that PhoP directly binds the identified PhoP box and that this site is important for the previously demonstrated regulation of OmpT by PhoP.

Omptin orthologues of E. coli exhibit differential activity toward substrates.

Although all are regulated by the same transcriptional regulator, it is possible that observed differences in activity could result from different degrees of induction due to promoter differences between omptin orthologues or due to altered recognition and processing of particular substrates. To distinguish between these, we created plasmid-based fusions in which we put the genes for each E. coli omptin orthologue (ompT, ompP, arlC, and croP) under the control of the ompT promoter in order to study the activity of the individual proteins alone, independent of cognate promoter strength. These fusions were then transformed to BW25113 ΔompT, and we assessed activity against a synthetic FRET substrate and also assessed the resistance of these strains to LL-37 and mCRAMP. As shown in Fig. 5A, we observed that despite being controlled by the same promoter, there were significant differences in the activity of each fusion toward this substrate. We also observed significant differences in the protection these omptins confer to host defense peptides human cathelicidin LL-37 (Fig. 5B) and murine mCRAMP (Fig. 5C). Expression of omptin genes ompT and arlC restored at least wild-type levels of resistance to LL-37. CroP conferred significantly increased resistance to mCRAMP, but not to LL-37, with the opposite being true for OmpT. ArlC, in particular, provided significantly increased resistance to both peptides. Furthermore, the same strains expressing the four different omptins differed in their ability to cleave an LL-37-based substrate assessed by FRET assay. We reasoned that although these omptins are expressed under the control of the same promoter, and therefore should be expressed at similar levels, posttranslational processing or altered protein stability might also play a role in the final levels of activity. We added an HA epitope tag to the C terminus of each of these omptins and detected protein levels by Western blotting. We observed different levels of protein as well as evidence of differential stability in each of the omptins studied. As described previously, omptin proteases appear to undergo autoproteolysis, and this appeared to occur differentially between omptins, likely resulting in different levels of functional omptin protease expressed by our plasmids (Fig. 5D). Although we didn't precisely control the levels of expressed omptin, we observed that in spite of having almost undetectable levels of protein, OmpP exhibited the highest protease activity against the LL-37 mimetic substrate while failing to provide significant protection from either LL-37 or mCRAMP. This is consistent with there being significant differences in substrate specificity unique to the omptins themselves and that the observed differences are not solely due to differences in expression.
FIG 5 Omptins exhibit differential cleavage activity of a host defense peptide-based probe and provide various levels of protection against host defense peptides. (A) Förster resonance energy transfer (FRET)-based activity of BW25113 ΔompT strains transformed with different omptins under the control of the ompT promoter. Cultures were grown to mid-log phase in M9 medium containing 100 μM Mg2+. (B and C) Percentages of survival of BW25113 ΔompT strains transformed with different omptins under the control of the ompT promoter after a 10-min exposure to 10 μg/mL of either (B) LL-37 or (C) mCRAMP. Cultures were grown to mid-log phase in M9 medium containing 100 μM Mg2+. Statistics indicate P values from a Kruskal-Wallis one-way ANOVA with Dunn’s multiple-comparison testing. Omptin-expressing strains were compared to the nonexpressing BW25113 ΔompT strain. Outliers were identified and removed using the ROUT method with Q set to 1%. ns, no statistical difference; ***, P < 0.001; ****, P < 0.0001. Statistics indicate P values from one-way ANOVA with Tukey’s multiple-comparison testing. Omptin-expressing strains were compared to the nonexpressing BW25113 ΔompT strain. ns, no statistical difference; *, P < 0.05; ****, P < 0.0001. (D) Western blotting for HA-tagged omptins expressed under the ompT promoter. Cultures were grown to mid-log in M9 medium containing 100 μM Mg2+. The results shown are the mean and standard error of the values represented in each treatment. ##, P < 0.01 when the DompT strain was compared to WT by Tukey's post-hoc test.
Based on previous reports of the role of omptin proteases in complement resistance (36), we also assessed the ability of these omptins to cleave complement components C3 and factor B (Fig. 6A and B). We saw that the omptins differed from each other in their abilities to cleave a given complement component but also that a given omptin could cleave one component better than the other. We observed that the croP-expressing strain exhibited the greatest production of cleaved C3α, while no cleavage was observed by the ompP-expressing strain (Fig. 6A). ArlC and OmpT exhibited moderate ability to cleave C3. In contrast, the ompT-expressing strain was unable to cleave factor B at all, and the most cleaved component, factor B, was detected after reactions with the croP- and arlC-expressing strains (Fig. 6B). We then assessed susceptibility of these strains to killing by normal human serum (Fig. 6C). We observed that the omptins that were able to cleave both products, ArlC and CroP, conferred the highest levels of resistance to serum. Taken together, these results demonstrate the variation in these omptins’ ability to cleave different substrates and show that these differences are important in the level of resistance they provide to innate immune responses, such as host defense peptides and complement (Table 1).
FIG 6 Omptins vary in their ability to cleave complement components and protect from killing by human serum. (A and B) Western blot analysis of cleavage of C3 (A) or factor B (B) protein by BW25113 ΔompT strains transformed with different omptins under the control of the ompT promoter. Mid-logarithmic cultures grown in M9 medium containing 100 μM Mg2+ were incubated with C3 or factor B protein for 2 h. (C) Percentage of survival of BW25113 ΔompT strains transformed with different omptins under the control of the ompT promoter after a 10-min exposure to human serum. Cultures were grown to mid-log phase in M9 medium containing 100 μM Mg2+. Statistics indicate P values from a Kruskal-Wallis one-way ANOVA with Dunn’s multiple-comparison testing. Omptin-expressing strains were compared to the nonexpressing BW25113 ΔompT strain. Outliers were identified and removed using the ROUT method with Q set to 1%. ns, no statistical difference; **, P < 0.01; ****, P < 0.0001.
TABLE 1 Summary of omptin activity toward the substrates tested
Omptin proteaseActivity toward:
LL-37mCRAMPHuman serumC3Factor B


The PhoPQ system is a well-characterized regulator of outer membrane susceptibility to cationic antimicrobial peptides (37). Much of this resistance occurs via modifications to the structure of lipopolysaccharide (LPS) that alter the surface charge and hydrophobicity of the molecule, leading to reduced interaction with cationic antimicrobial peptides. In addition to this LPS modification-mediated antimicrobial peptide resistance, Enterobacterales also produce omptin proteases that can target components of the host innate immune system. Here, we demonstrate that omptin proteases are another conserved antimicrobial peptide resistance mechanism that is regulated by the PhoPQ system.
In E. coli, at least three omptin proteases have been identified to date: the housekeeping chromosomally encoded OmpT (38), the F′ episome-encoded OmpP (39), and a plasmid-encoded protein, ArlC (24). Some strains of E. coli, like APECO1 (40), also carry a second chromosomal copy of the ompT protease, although the consequences of this increased copy number are unknown. Citrobacter rodentium encodes the OmpT homologue CroP (41). Although these proteins have been associated with in vivo fitness and virulence, a detailed examination of their regulation has not been undertaken. There are previous reports suggesting a role of the PhoPQ system in regulation of omptins (4244). MicA, a small RNA (sRNA) repressor of phoP, represses ompT transcription (45). Consistent with these previous reports, our findings in E. coli BW25113 show that OmpT activity, expression, and protein levels are induced by PhoPQ-inducing conditions of low Mg2+ and are absent when phoP is deleted (Fig. 1). We also identified a canonical PhoP-binding site in the promoters of these omptins and show that mutation of this site abrogates PhoP-dependent regulation of ompT expression (Fig. 4A to C). Furthermore, we observed that mutation of this site interferes with PhoP binding (Fig. 4D), clearly demonstrating that PhoP regulates omptins by directly binding to this site on the promoter of these omptin genes.
Omptin activity in C. rodentium and a number of E. coli strains is dependent on the presence of phoP (Fig. 2). The E. coli strains tested carry different combinations of OmpT, OmpP, ArlC, and CroP, suggesting that PhoP-dependent omptin regulation is not limited to OmpT. Indeed, we observed that expression of ompP, arlC, and croP is also absent when PhoP is repressed or deleted (Fig. 3). We also identified similar putative PhoP-binding sites in the promoters of these omptin genes, suggesting a conserved mechanism among these omptins.
The observation that omptin regulation by PhoP is conserved in OmpT, OmpP, ArlC, and CroP suggests that each allele might provide substrate selectivity favoring the maintenance of one allele or another, depending upon the strain and the environments in which that strain is found. Previous comparisons of OmpP and OmpT have identified some differences in the substrate specificity of the enzymes, although both were able to protect the expressing strains from protamine exposure (27). Similarly, previous work has shown that both OmpT and ArlC contribute to host defense peptide resistance, with significant differences in the resistance profile of isogenic strains lacking either ArlC or OmpT, suggesting that these proteins have different substrate specificities (24). Divergent omptin substrate selectivity has been observed previously between OmpT and CroP, in which OmpT preferentially cleaves antimicrobial peptides with α-helical structure, while CroP exhibits a preference for unfolded/unstructured peptides (23). Similarly, the Shigella virulence-related omptin protein SopA has lost generic broad-substrate activity, appears to have become tightly tuned to a single substrate (IcsA), and maintains this substrate with a strict polar localization (46, 47). The mechanisms for omptin substrate selectivity are not well characterized. Previous work comparing OmpP to OmpT identified a single residue near the active site of OmpP (V214) that, when mutated to the analogous OmpT site aspartate, changed the activity of the resultant mutant to an OmpP substrate (27). The equivalent site in ArlC is asparagine, suggesting that this heterogeneity might contribute to the observed altered substrate selectivity (see Fig. S1 in the supplemental material).
Omptin proteases have a wide variety of substrates that may contribute to in vivo fitness of the bacterium containing them, including host defense peptides (HDPs) like LL-37 or mCRAMP (23, 24), components of the complement cascade or the clotting cascade (36, 48), and nonphysiological but cationic substrates like protamine (25). Our data show that OmpT, OmpP, ArlC, and CroP conferred different benefits to survival when parent strains expressing each of these omptins were confronted with HDPs or whole human serum (Fig. 5A and B and Fig. 6C). These differences in protective ability translated to differences in the ability of these omptins to cleave an HDP-based FRET substrate as well as factor B and C3, components of complement (Fig. 5C and Fig. 6A and B). Interestingly, while each omptin allele demonstrated differential activity toward the tested substrates, the ArlC protein showed high-level activity against all substrates. It was also the most protective against killing by LL-37, mCRAMP, and human serum, suggesting that this omptin (which is also only found on extrachromosomal plasmids) may be an important contributor to host resistance in E. coli strains that carry it.
The differences in activity and unique survival advantages provided by each omptin might be why specific omptins are retained or lost in different strains, even though they are regulated in the same manner. Alterations in either the activity of omptins or in their transcriptional activity have been associated with changes in virulence, including in pandemic Y. pestis or in invasive Salmonella enterica (49), suggesting that omptin proteases are critical factors for interaction with and resistance to components of the host innate immune system. The specific activity of the omptin possessed by a given bacterium is also known to be essential for its virulence program, such as Pla in Y. pestis, which cleaves plasminogen activator inhibitor 1 and is crucial for dissemination (48, 50), and the previously mentioned SopA is crucial for Shigella’s characteristic cell-to-cell spread (47). CroP is found in C. rodentium, a murine pathogen, and has been described as particularly efficient at cleaving mCRAMP, consistent with our results here (40). Even the loss of specific omptins can be advantageous, depending on the specific virulence program of a bacteria: all Shigella strains have lost OmpT, which inhibits intracellular spreading when present due to cleavage of the virulence-associated cell spreading factor IcsA (51).
Although we show that PhoP regulation of the E. coli omptins is conserved, there are likely other regulatory inputs that might differentially affect omptin protease activity. Alterations in the promoter sequence can affect the expression levels of the protein, with significant impacts on virulence. This has been demonstrated previously in enterohemorrhagic E. coli (EHEC), in which the ompT promoter in EHEC supports significantly increased expression of OmpT protein, with increased resistance to LL-37 (28), and a single nucleotide polymorphism in pandemic S. enterica affecting the activity of PgtE has also been described (49). In addition, other regulatory inputs could contribute to altered expression levels among the omptin orthologues that could alter activity of these proteases. This has been demonstrated previously for the PgtE protease of Salmonella, which is subject to significant silencing by Hns (52) and positive regulation by the SlyA and PhoP proteins (53) and by the CpxRA system (54). Even the prototypical OmpT protein shows evidence of regulation by small RNAs (55).
Beyond transcriptional regulation, the physiological state of the bacteria may have posttranslational regulatory effects on omptin activity. LPS is required for omptin activity of OmpT, Pla, and PgtE, with specific LPS structural requirements for proteolytic activity to occur (21, 56, 57). The length of O antigen can also affect omptin activity (57, 58). As environmental changes result in alterations to LPS structure, omptin activity may be affected in kind. The OmpT regions proposed to participate in LPS binding are slightly different in the other omptins OmpP, SopA, and PgtE (59), potentially changing how LPS modifications might affect protease activity in these omptins. This paints a very complex picture of omptin regulation. Nonetheless, our results suggest that regardless of such strain-to-strain differences affecting omptin activity, the activity of the E. coli and C. rodentium omptins is largely dependent on PhoP. Whether other environmental conditions or regulatory proteins might further contribute to differential omptin regulation remains an unanswered, but important, question.
Our observations come with several caveats. The first is that the omptins studied in their native host strains were under the control of their native promoter, and it is therefore difficult to distinguish between altered expression levels and altered substrate specificity. We addressed this by expressing each of the ompT, ompP, arlC, and croP genes from the ompT promoter and by constructing versions of these alleles containing an HA epitope tag to the C terminus to monitor protein levels. We then expressed each of these alleles in the omptin-deficient E. coli strain BL21. It is important to note that these epitope-tagged proteins are only useful for assessing production levels, due to the need for an unaltered C terminus to get efficient targeting to the outer membrane via the Bam complex (60). As seen in Fig. 5D, even having controlled for this aspect of expression, we observed differential proteolysis of the omptin proteins, suggesting that there are other factors that might contribute to the amount of functional protease expressed by our plasmids. With this in mind, however, we observed that the least stable proteins (i.e., ArlC and CroP) are the ones that provide the most protection from LL-37, mCRAMP, and human serum. This is consistent with the hypothesis that the differences in protein levels observed are not responsible for the differences in bacterial protection that we observed.
Understanding the mechanisms by which omptin proteases are regulated, as well as how they mediate substrate selectivity, may shed light on how virulence emerges in these microbes. Although OmpT appears to be conserved in the majority of E. coli strains, both OmpP and ArlC are associated with mobile genetic elements, suggesting that the acquisition of one of these proteins could open up new niches for colonization or invasion by these microbes. Here, we provide evidence that although the omptins of E. coli are regulated in a conserved manner, they still exhibit significant differences in behavior—supporting the hypothesis that they are associated with altered virulence or host adaptation.


Bacterial strains and growth conditions.

All strains used in this study are shown in Table 2. Bacteria were routinely grown at 37°C with shaking in lysogeny broth (LB) for culturing and for molecular manipulations. For antibiotic selection, we routinely used ampicillin at 100 μg/mL or kanamycin at 50 μg/mL. For the defined medium composition, we used a modified N-minimal (61) or M9 medium. N-minimal medium contained 5 mM KCl, 7.5 mM (NH4)2SO4, 0.5 mM K2SO4, 1 mM KH2PO4, 0.1 mM Tris-HCl (pH 7.4), 10 mM or 10 μM MgCl2, 1 μM or 100 μM Fe3+, and 0.2% glucose. M9 medium contained 1 × M9 salts supplemented with 0.1% Casamino Acids, 0.001 mg/mL thiamine, 0.01 mM FeSO4, 0.1 mM CaCl2, 0.1 mM MgSO4, and 0.4% (wt/vol) glucose.
TABLE 2 Escherichia coli and Citrobacter rodentium strains used in this study
Bacterial strainCharacteristicaSource
E. coli  
 BW25113Parent strain of Keio collectionBaba et al., 2006 (64)
 BW25113 ΔphoPphoP mutant of BW25113Baba et al., 2006 (64)
 BW25113 ΔpmrApmrA mutant of BW25113Baba et al., 2006 (64)
 NRG857cAIECEaves-Pyle et al., 2008 (31)
 NRG857c ΔphoPphoP mutant of AIECMcPhee et al., 2014 (24)
 LF82AIECBoudeau et al., 1999 (30)
 LF82 ΔphoPphoP mutant of AIECThis study
 EHEC 86-24EHECGriffin et al., 1988 (32)
 EHEC 86-24 ΔphoPphoP mutant of EHECKunwar et al., 2020 (65)
C. rodentium  
 DBS100WT C. rodentiumSchauer et al., 1995 (66)
 DBS100 ΔphoPQphoPQ mutant of C. rodentiumReid-Yu et al., 2015 (33)
AIEC, adherent-invasive E. coli; EHEC, enterohemorrhagic E. coli.


All plasmids and primers used in this study are shown in Table 3 and Table 4. Chemically competent cells were prepared as previously described (62). The ompT promoter-omptin fusions were created using splicing by overlap extension PCR (SOE-PCR) to create fusions between the promoter of chromosomal ompT with the coding regions of ompT, ompP, or arlC. These PCR products were then cloned into pCR2.1-TOPO according to the manufacturer’s instructions and then subcloned into pWSK129 using XbaI and KpnI. These constructs were then transformed to BW25113 ΔompT, and the activities of the constructs were assessed by the FRET assay as described below. We similarly used SOE-PCR to create translational reporter fusion between the promoters of ompT, ompP, arlC, or croP and the GFP-mut3 gene, where the GFP reporter replaced the entire coding region of the omptin gene itself, and these were cloned into pCR2.1-TOPO. For the construction of the pCR-ompT-HSmut-GFP PhoP-binding site mutant plasmid, we carried out site-directed mutagenesis of pCR-ompT-GFP according to the Quikchange II site-directed mutagenesis kit (Agilent Technologies). These were transformed into BW25113, BW25113 ΔphoP, and BW25113 ΔpmrA, and transcription was measured by GFP assay as described below.
TABLE 3 Plasmids used in this study
pCR2.1-TOPO Invitrogen
pCR2.1-ompT-GFPGFP fusion to ompTThis study
pCR-ompP-GFPGFP fusion to ompPThis study
pCR-arlC-GFPGFP fusion to arlCThis study
pCR-croP-GFPGFP fusion to croPThis study
pCR-pompT-ompTompT expressed under control of ompT promoterThis study
pCR-ompT-HSmut-GFPompT expressed under control of mutated ompT promoterThis study
pWSK129-pompT-ompTompT expressed under control of ompT promoterThis study
pWSK129-pompT-ompPompP expressed under control of ompT promoterThis study
pWSK129-pompT-arlCarlC expressed under control of ompT promoterThis study
pWSK129-pompT-croPcroP expressed under control of ompT promoterThis study
pWSK129-phoP-HAHA-tagged phoPThis study
pWSK129-pompT-ompT-HAHA-tagged ompT expressed under control of ompT promoterThis study
pWSK129-pompT-ompP-HAHA-tagged ompP expressed under control of ompT promoterThis study
pWSK129-pompT-arlC-HAHA-tagged arlC expressed under control of ompT promoterThis study
pWSK129-pompT-croP-HAHA-tagged croP expressed under control of ompT promoterThis study
TABLE 4 Primers used in this study
PrimerSequence (5′→3′)aDescriptionb
ompT-ompT P1GGTACCGACGGGCATAAATGAGGAAGExpression of omptins under ompT promoter
ompT-ompT-P4TCTAGATTAAATGTGTACTTAAGACCExpression of ompT under ompT promoter
ompT-ompP-P4TCTAGATTAAAACGTGTACTTCAGACCExpression of ompP under ompT promoter
ompT-croP-P4TCTAGATCAGAAGGTATATTTGAGGCCExpression of croP under ompT promoter
ompT-arlC-P4TATAGATTAAAATAATACTTCAGACCExpression of arlC under ompT promoter
ompT-GFP P1GACGGGCATAAATGAGGAAGExpression of GFP under ompT promoter
ompT-GFP P6GGGATGAAGGAACGTCATTTACExpression of GFP under ompT promoter
ompP-GFP P1GGATCCACTGAGGACTTTATCCAGGCExpression of GFP under ompP promoter
arlC-GFP P1GGATCCTTTCGTCGAATGGTAATGTCExpression of GFP under arlC promoter
croP-GFP P1GGATCCGCTGGCCGGATGGTATAAAGExpression of GFP under croP promoter
omptin-GFP P4CTGCAGTTATTTGTATAGTTCATCCATExpression of GFP under omptin promoter
pmrDprom-FTGCATTATCCTGTTTGCTAAGAGTBiotin-tagged promoter probes for pulldown assay
pmrDprom-R biotinBtn-CCACGAGAGCGTTGATCTTAGTGBiotin-tagged promoter probes for pulldown assay
ompTprom-FAAACCTCATGCTATTTTCGCTTABiotin-tagged promoter probes for pulldown assay
ompTprom-R biotinBtn-AAAAGTTCTCCATTCAATCGTTTTBiotin-tagged promoter probes for pulldown assay
dnaKprom-FAGCCGCTATCGAAGCGAAAATGCBiotin-tagged promoter probes for pulldown assay
dnaKprom-R biotinBtn-CTTTTTTAAATTGCCCCTAGATGABiotin-tagged promoter probes for pulldown assay
Btn, biotin.
HS, half-site.

FRET-based omptin protease assay.

We developed a probe in which an internal 11-amino-acid sequence of the human peptide LL-37 (LLGDFFRKSKEKIGKEFKRIVQRIKDFLRNLVPRTES) was labeled on the N terminus with a fluorescent probe, while the C terminus was tagged with a proprietary quenching molecule (5-carboxyfluorescein [5-FAM]–GKEFKRIVQRI–K-QXL520). When this molecule is cleaved, quenching is relieved and fluorescence can be monitored. Strains were grown overnight in M9 or N-minimal medium at 37°C with shaking. The overnight culture was washed twice, diluted 1:50 in fresh N-minimal medium containing either low Mg2+ and low Fe3+ (10 μM Mg2+ and 1 μM Fe3+), high Mg2+ and low Fe3+ (10 mM Mg2+ and 1 μM Fe3+), or high Mg2+ and high Fe3+ (10 mM Mg2+ and 100 μM Fe3+) and grown to mid-log phase at 37°C. The cultures were washed twice in 10 mM HEPES buffer (pH 7.2) and then normalized to an optical density at 600 nm (OD600) of 0.5. The FRET substrate was added to a final concentration of 7 μg/mL to ~1.1 × 107 cells or HEPES buffer as a blank. Fluorescence was measured over 1 h at 25°C with an excitation wavelength of 485 nm and emission wavelength of 528 nm. The initial slope of the activity was calculated, and the measurements shown represent the average of at least three independent experiments.

Translational reporter assay (GFP assay).

Overnight cultures were grown in LB plus 20 mM MgSO4. The next day, the cultures were washed twice in 1× N-minimal salts and normalized to an OD600 of ~0.02 in N-minimal medium containing either low Mg2+ and low Fe3+ (10 μM Mg2+ and 1 μM Fe3+), high Mg2+ and low Fe3+ (10 mM Mg2+ and 1 μM Fe3+), or high Mg2+ and high Fe3+ (10 mM Mg2+ and 100 μM Fe3+). These cultures were added to black, clear-bottom 96-well culture plates and measured every 30 min for fluorescence (excitation at 485 nm and emission at 530 nm) and absorbance (600 nm). The ratio of fluorescence to absorbance was calculated at a mid-logarithmic growth point, and the ratios are reported as relative fluorescence unit (RFU)/OD600.

Western blotting.

Bacteria were grown with shaking in LB at 37°C overnight. Cells were then subcultured in N-minimal medium containing either low Mg2+ and low Fe3+ (10 μM Mg2+ and 1 μM Fe3+), high Mg2+ and low Fe3+ (10 mM Mg2+ and 1 μM Fe3+), or high Mg2+ and high Fe3+ (10 mM Mg2+ and 100 μM Fe3+) until the early stationary phase was reached. The cells obtained were normalized to an OD600 of 1 (2 × 109 cells) and washed once in prechilled phosphate-buffered saline (PBS). The pellets were resuspended in 1× DNase I reaction buffer (10 mM Tris-HCl, 50 mM NaCl, and 10 mM MgCl2 at pH 7.5) and subjected to lysis via three freeze-thaw cycles prior to incubation with 10 mg/mL DNase I at 37°C for 1 h. Samples were incubated at 100°C for 10 min in 1× bicinchoninic acid (BCA)-compatible lysis buffer (4% SDS and 0.2 M Tris-HCl [pH 6.8]). The otal protein concentration in each sample was measured using a Pierce BCA assay kit (Thermo Fisher, Inc.), and samples were mixed with postlysis Laemmli buffer (2% SDS, 60% glycerol, 1.2 M dithiothreitol [DTT], and 0.1 M Tris-HCl [pH 6.8]). Proteins were resolved by SDS-PAGE on a 6% stacking gel and 12% separating gel, followed by electroblotting to nitrocellulose membranes at 200 V for 1 h. Membranes were blocked with 5% skim milk powder dissolved in 20 mM Tris–150 mM NaCl (pH 7.4) containing 0.1% Tween 20 (TBST) overnight at 4°C. They were then washed 3 times in TBST before incubation in rabbit anti-OmpT (Flare Biotech LLC) or mouse anti-DnaK (Enzo Life Sciences) for 1 h at room temperature, followed by being washed 3 times again in TBST and incubation in horseradish peroxidase (HRP)-conjugated goat anti-rabbit (Cell Signaling Technology) or HRP-conjugated goat anti-mouse (Promega) antibody for 1 h at room temperature. Following a third round of TBST washes, ECL (enhanced chemiluminescence) substrates A (1.5 mM Tris-HCl [pH 8.8], 90 mM coumaric acid, and 250 mM luminol) and B (30% H2O2 in distilled water [dH2O]) were mixed in a 1,000:3 ratio and added to the blots prior to visualization. All reagents used in these experiments were obtained from BioShop Canada unless otherwise indicated. Images were analyzed using ImageJ.

PhoP-HA pulldown assays.

Pulldown assays were conducted as described previously with modifications (63). Briefly, strains expressing PhoP-HA grown with shaking in LB at 37°C overnight. Cells were then subcultured in N-minimal medium with 10 μM Mg2+. Following two washes in PBS, cells were lysed by sonication (20 s at 10% amplitude with a 1-min rest for 10 cycles). Lysed bacteria were centrifuged for 20 min at 13,000 × g at 4°C. Protein content was quantified in this lysate by BCA assay. Biotin-tagged probes were synthesized by PCR using the primers in Table 4 and purified using a standard column clean kit, and the concentration was quantified by measuring absorbance at 260 nm. We mixed 5 μg of biotin-tagged promoter probes with 40 μL streptavidin-agarose bead slurry (20-μL packed bead volume) (Millipore Sigma) and topped up to 400 μL with BS/THES buffer (described below). This was incubated for 30 min at room temperature with rocking. Following washing 3 times in BS/THES buffer (22 mM Tris-Cl [pH 7.5], 4.4 mM EDTA, 8.9% [wt/vol] sucrose, 62 mM NaCl, 0.04% phosphatase inhibitor, 10 mM HEPES, 5 mM CaCl2, 50 mM KCl, and 12% glycerol) with 10 μg/mL salmon sperm DNA, 1,000 μg lysate was added, and the reaction mixture was topped up with BS/THES buffer with 20 μg/mL salmon sperm DNA and cOmplete protease inhibitor (Sigma-Aldrich) to a total volume of 500 μL. The bead-promoter-PhoP-HA complexes were pulled down by centrifugation at 500 × g for 5 min and washed in fresh BS/THES buffer with 10 μg/mL salmon sperm DNA and protease inhibitor. The wash step was repeated 3 times. Laemmli buffer was added, and samples were boiled at 100°C for 10 min. Proteins were visualized by Western blotting as described above. Rabbit anti-HA antibodies (Life Technologies) and HRP-conjugated goat anti-rabbit antibodies (Cell Signaling Technologies) were used for visualization.

Immunoblotting for C3 and factor B cleavage by omptin proteases.

Bacteria were grown with shaking in LB at 37°C overnight. Cells were then subcultured in M9 medium and washed in 10 mM HEPES (pH 7.2). Cultures were normalized to 1 × 108 CFU/mL. Bacteria were exposed to either human C3 or human factor B protein (Complement Technology, Inc.) in a 1:3 ratio and incubated for 2 h at 37°C with shaking. Following treatment, Laemmli buffer was added to the incubated mixture (1:1) and samples were boiled at 100°C for 5 min. Proteins were resolved by SDS-PAGE, transferred to nitrocellulose membrane, and developed according to the Western blot protocol described above. Proteins were visualized using primary antibodies (goat anti-human C3 or goat anti-human factor B antibodies; Complement Technology, Inc.) and secondary antibodies (HRP-conjugated rabbit anti-goat antibodies; Jackson ImmunoResearch Laboratories).

HDP killing assay.

Strains were grown overnight in minimal M9 medium at 37°C with shaking. The overnight culture was diluted 1:20 in fresh M9 medium and grown to mid-log phase at 37°C with shaking. The culture was then washed in 10 mM HEPES buffer (pH 7.2) and normalized to 108 CFU/mL. This normalized culture was treated with an equal volume of 1× PBS buffer or 100 μg/mL LL-37 peptide (for a treatment concentration of 50 μg/mL LL-37). A sample of the reaction mixture was taken at the time of treatment and 10 min following treatment. These samples were serially diluted in PBS and plated onto LB agar plates. Survival percentages were calculated by comparing the surviving bacteria at t = 10 min to the number of colonies at t = 0 min and normalized to the blank (PBS treatment). For each strain, 4 to 6 replicates were performed to obtain a mean percentage of survival. The HBD3 screen was performed the same way, using a peptide concentration of 2 μg/mL for a treatment concentration of 1 μg/mL. Survival percentages were log transformed before being plotted with GraphPad Prism 9 software.

Complement killing assay.

Strains were grown overnight in minimal M9 medium at 37°C with shaking. The overnight culture was diluted 1:20 in fresh M9 medium and grown to mid-log phase at 37°C with shaking. The culture was then washed in 10 mM HEPES buffer (pH 7.2) and normalized to 107 CFU/mL. The normalized culture was treated with a 3:1 amount of heat-inactivated human serum or normal human serum for 10, 30, and 60 min. Sterile-filtered human male AB plasma was used in this experiment (Sigma-Aldrich, St. Louis, MO, USA). Following treatment, the reaction mixtures were serially diluted in PBS and plated onto LB agar plates. Survival percentages were calculated by comparing the surviving bacteria at t = 10 min to the number of colonies at t = 0 min and normalized to the blank (inactivated serum treatment).


This work was supported by a National Sciences and Engineering Research Council Discovery Grant (RGPIN-04679-2015) and by startup funding from the Ryerson University Faculty of Science to J.B.M.

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Published In

cover image Infection and Immunity
Infection and Immunity
Volume 91Number 124 January 2023
eLocator: e00518-22
Editor: Andreas J. Bäumler, University of California, Davis
PubMed: 36533918


Received: 24 November 2022
Accepted: 30 November 2022
Published online: 19 December 2022


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  1. Escherichia coli
  2. omptin proteases
  3. two-component regulation
  4. complement resistance
  5. host-defense peptide resistance
  6. PhoPQ
  7. antimicrobial peptides
  8. ompT
  9. omptins
  10. two-component regulatory systems



Youn Hee Cho
Department of Chemistry and Biology, Toronto Metropolitan University, Toronto, Ontario, Canada
Monir Riasad Fadle Aziz
Department of Chemistry and Biology, Toronto Metropolitan University, Toronto, Ontario, Canada
Avery Malpass
Department of Chemistry and Biology, Toronto Metropolitan University, Toronto, Ontario, Canada
St. Francis Xavier University, Antigonish, Nova Scotia, Canada
Tanuja Sutradhar
Department of Chemistry and Biology, Toronto Metropolitan University, Toronto, Ontario, Canada
Jasika Bashal
Department of Chemistry and Biology, Toronto Metropolitan University, Toronto, Ontario, Canada
Veronica Cojocari
Department of Chemistry and Biology, Toronto Metropolitan University, Toronto, Ontario, Canada
Department of Chemistry and Biology, Toronto Metropolitan University, Toronto, Ontario, Canada


Andreas J. Bäumler
University of California, Davis


The authors declare no conflict of interest.

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