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Research Article
1 October 2019

Transcription of the Subtilase Cytotoxin Gene subAB1 in Shiga Toxin-Producing Escherichia coli Is Dependent on hfq and hns


Certain foodborne Shiga toxin-producing Escherichia coli (STEC) strains carry genes encoding the subtilase cytotoxin (SubAB). Although the mode of action of SubAB is under intensive investigation, information about the regulation of subAB gene expression is currently not available. In this study, we investigated the regulation of the chromosomal subAB1 gene in laboratory E. coli strain DH5α and STEC O113:H21 strain TS18/08 using a luciferase reporter gene assay. Special emphasis was given to the role of the global regulatory protein genes hfq and hns in subAB1 promoter activity. Subsequently, quantitative real-time PCR was performed to analyze the expression of Shiga toxin 2a (Stx2a), SubAB1, and cytolethal distending toxin V (Cdt-V) genes in STEC strain TS18/08 and its isogenic hfq and hns deletion mutants. The deletion of hfq led to a significant increase of up to 2-fold in subAB1 expression, especially in the late growth phase, in both strains. However, deletion of hns showed different effects on the promoter activity during the early and late exponential growth phases in both strains. Furthermore, upregulation of stx2a and cdt-V was demonstrated in hfq and hns deletion mutants in TS18/08. These data showed that the expression of subAB1, stx2a, and cdt-V is integrated in the regulatory network of global regulators Hfq and H-NS in Escherichia coli.
IMPORTANCE Shiga toxin-producing Escherichia coli (STEC) strains are responsible for outbreaks of foodborne diseases, such as hemorrhagic colitis and the hemolytic uremic syndrome. The pathogenicity of those strains can be attributed to, among other factors, the production of toxins. Recently, the subtilase cytotoxin was detected in locus of enterocyte effacement (LEE)-negative STEC, and it was confirmed that it contributes to the cytotoxicity of those STEC strains. Although the mode of action of SubAB1 is under intensive investigation, the regulation of gene expression is currently not known. The global regulatory proteins H-NS and Hfq have impact on many cellular processes and have been described to regulate virulence factors as well. Here, we investigate the role of hns and hfq in expression of subAB1 as well as stx2a and cdt-V in an E. coli laboratory strain as well as in wild-type STEC strain TS18/08.


Shiga toxin-producing Escherichia coli (STEC) strains are a major cause of serious foodborne diseases, leading to diarrhea, hemorrhagic colitis, and the hemolytic uremic syndrome (HUS) (1, 2). The main virulence factors described so far are the production of Shiga toxins (Stx) (3) and the components of the locus of enterocyte effacement (LEE) (4). Although the LEE is widely distributed in STEC, many LEE-negative STEC strains have been isolated and characterized (5, 6). Characterization of those strains led to identification of other putative virulence factors, such as the subtilase cytotoxin (SubAB). So far, SubAB was found to be produced only by LEE-negative STEC strains and was first identified in E. coli O113:H21 strain 98NK2, involved in an HUS outbreak (7). SubAB belongs to the family of AB5 toxins and consists of a catalytic A subunit and a homopentameric B subunit, the latter mediating binding to the terminal sialic acids of glycoproteins of the host cell (8). The catalytic activity of SubA is the cleavage of the endoplasmic chaperone BiP/GRP78, leading to an unfolded protein response and apoptosis (9, 10).
Recent studies demonstrated the contribution of SubAB to STEC cytotoxicity (11, 12). Until now, three genetic subAB variants have been described in STEC strains. Whereas subAB1 is located on the virulence plasmid pO113, the variants subAB2-1, subAB2-2, and subAB2-3 are located on the chromosome (13, 14). The regulatory pathways leading to increased expression of subAB are currently not known.
Global regulatory proteins regulate many cellular processes in bacteria. The host factor of bacteriophage Qβ (Hfq) is an RNA chaperone and interacts with mRNA and small regulatory RNAs in E. coli (15). Hfq itself or the RNAs that are processed by Hfq are implicated in regulation of metabolism, quorum sensing, and regulation of adhesion and motility (16, 17). Moreover, Hfq regulates many stress responses, such as response to acids or oxidative stress (18, 19). In addition to Hfq, the histone nucleoid structuring protein H-NS is a pleiotropic regulator. It regulates transcription by binding directly to DNA or to the RNA polymerase (20). H-NS is known to alter DNA topology (20) and to regulate many processes during the bacterial life cycle. It regulates metabolic processes and is integrated in the regulatory pathway of genes involved in motility (21). Moreover, H-NS plays a role in stress response, such as cold shock response (22). Regulation of cellular processes by Hfq and H-NS have been studied extensively in recent years. Both Hfq and H-NS are integrated in the regulation of virulence factors of pathogenic bacteria as well. Kendall et al. identified the role of Hfq in regulation of virulence in enterohemorrhagic E. coli (EHEC) O157:H7. In this study, it was shown that deletion of the hfq gene led to an increase of stx2AB expression (23). The nucleoid binding protein H-NS silences foreign DNA with high percentages of adenosine and thymine (A/T) content (24) and is therefore integrated in regulation of virulence genes. H-NS was also identified to regulate genes of the LEE. H-NS also regulates expression of genes of the hemolysin in EHEC (25).
First insights in gene expression of subAB1 were given by Hauser et al. (12). In STEC strain TS18/08, isolated from minced meat, the expression of subAB1 was analyzed using quantitative real-time PCR (qRT-PCR). It was shown that the expression of subAB1 was highest in the exponential growth phase under standard batch culture conditions (12).
The aim of this study was to investigate whether subAB1 expression is under the control of Hfq and H-NS. Therefore, deletion of hfq and hns genes was performed, and the expression of subAB1 was measured with a luciferase reporter gene assay in laboratory E. coli strain DH5α and the foodborne STEC strain TS18/08 (26). E. coli TS18/08 produces three toxins: the subtilase cytotoxin SubAB1, Shiga toxin 2a (Stx2a), and the cytolethal distending toxin V (Cdt-V) (11). Therefore, we were interested in whether the expression of the corresponding toxin genes is dependent on the global regulators Hfq and H-NS. Quantitative real-time PCR was applied to study in vitro transcription of those toxin genes in E. coli TS18/08.


Transcription of cloned hfq or hns genes present on complementation plasmids.

Prior to the use of plasmids pLH01 and pLH02 for complementation studies, expression of cloned hfq and hns genes on these plasmids was investigated. Therefore, deletion mutants DH5α Δhfq/pHL01 and DH5α Δhns/pHL02 were grown under standard batch conditions and total RNA was isolated. Transcription analyses of hfq and hns were conducted as described previously. PCR analysis with hfq- and hns-specific primers and subsequent agarose gel electrophoresis of the resulting cDNA demonstrated PCR products with the molecular size of hfq (309 bp) and hns (414 bp). Laboratory strain E. coli DH5α (positive control), as well as its E. coli DH5α Δhfq and DH5α Δhns (negative controls) isogenic deletion mutants, were also applied in the procedure. cDNAs of the hfq gene and hns gene were not amplified in either control, as representatively shown for the E. coli DH5α Δhfq strain (see Fig. S1 in the supplemental material). For both the wild-type strain and the E. coli DH5α Δhfq/pLH01 and DH5α Δhns/pLH02 complemented strains, bands with an expected size of around 300 bp and 414 bp, respectively, were visible on the agarose gels.
Reverse transcriptase negative controls show that no cross-contamination with DNA occurred during the experimental procedure. This indicated that the plasmids pLH01 and pLH02 expressed the genes of interest and can be used for complementation of deletion mutants.

Deletion of hfq and hns leads to slower growth in STEC TS18/08.

In order to investigate the effects of hfq and hns on subAB gene expression, growth curves of E. coli DH5α (Fig. 1) and STEC strain E. coli TS18/08 (Fig. 2), as well as of their respective hfq and hns mutants, were initially compared. It could be shown that deletion of either hfq or hns led to a minor decrease in growth in DH5α and both mutants, whereas the highest decreases in values of optical density at 600 nm (OD600) were measured as 1.6-fold in the DH5α Δhns strain and 1.8-fold in the DH5α Δhfq strain in the late growth phase. This effect was stronger in STEC TS18/08 than in DH5α. The deletion mutants showed significantly lower OD600 values over the cultivation period. Whereas deletion of hns led to a decrease of up to 4.6-fold in the exponential growth phase, the decline in OD600 values was 2.3-fold lower in the TS18/08 Δhfq strain than in the wild type. After 5 h of incubation, decrease rates diminished up to 1.5-fold and 1.3-fold in TS18/08 Δhns and TS18/08 Δhfq strains, respectively. The reduction in OD600 values in the TS18/08 Δhns strain could be restored after complementation with the wild-type gene on plasmid pLH02 (Fig. 2). However, reduction in OD600 values could not be restored in the TS18/08 Δhfq strain by complementation with plasmid pLH01. Although TS18/08 Δhfq and TS18/08 Δhns deletion mutants grew more slowly than wild-type TS18/08, all strains reached the exponential growth phase in a period of 5 h. Therefore, a growth period spanning 5 h was chosen for further investigations of PsubAB1 activity.
FIG 1 Growth curves of E. coli DH5α wild-type and mutant strains. Cultivation was conducted for 5 h in LB at 37°C with aeration. The OD600 was determined over a period of 5 h for E. coli DH5α/pKMD3, DH5α Δhns/pKMD3, and DH5α Δhfq/pKMD3 strains. Data represent mean values and standard deviations from three biological replicates.
FIG 2 Impact of deletion of genes hfq and hns on growth of STEC O113:H21 strain TS18/08. Optical density of wild-type strain (TS18/08), TS18/08/pKMD3 + pBR322 (TS18/08/pBR322) empty vector backbone control, TS18/08 Δhns/pKMD3 (TS18/08 Δhns) and TS18/08 Δhfq/pKMD3 (TS18/08 Δhfq) deletion mutants, and TS18/08 Δhns/pKMD3 + pLH02 (TS18/08 Δhns + hns) and TS18/08 Δhfq/pKMD3 + pLH01 (TS18/08 Δhfq + hfq) complemented strains is shown for measurements over 5 h. Data from three biological replicates are shown; error bars indicate standard deviations from mean values. Asterisks indicate statistical significance (P < 0.05) compared to values for TS18/08.

E. coli DH5α/pKMD3 is suitable for measuring subAB1 promoter activity.

Promoter activity of subAB1 was measured as described above in E. coli DH5α, E. coli TS18/08, and their respective Δhfq and Δhns mutants. As shown in Fig. 3, the relative promoter activity of subAB1 (PsubAB1) of E. coli DH5α/pKMD3 significantly increased up to 11-fold compared to the starting value in the exponential growth phase, showing the highest activity of 2.29 × 105 relative light units (RLU)/OD600 after 2 h of cultivation. This activity did not increase any further. Similar to the activity measured in E. coli DH5α/pKMD3, the PsubAB1 activity of TS18/08/pKMD3 increased significantly up to 7-fold compared to the starting value in the exponential growth phase and did not increase further. The highest PsubAB1 activity in STEC TS18/08 was detected with 3.3 × 105 RLU/OD600 after 3 h of cultivation (Fig. 3). The results of these experiments demonstrated that the luciferase reporter system worked well for measuring subAB1 promoter activity.
FIG 3 Comparison of PsubAB1 promoter activities in DH5α and TS18/08. Relative reporter gene activity (RLU/OD600) is shown. Bars represent data for E. coli DH5α (dark blue) and an STEC strain (TS18/08, light blue). OD600 is shown as squares. Cultivation was conducted for 5 h in LB at 37°C with aeration. Data from three biological replicates are shown; error bars indicate standard deviations from mean values.

Hfq represses PsubAB1 activity in E. coli DH5α and TS18/08 in the late exponential growth phase.

In Fig. 4, the effect of the hfq deletion in E. coli DH5α (Fig. 4A) and E. coli TS18/08 (Fig. 4B) on PsubAB1 activity is shown. In the E. coli DH5α Δhfq/pKMD3 mutant, the relative reporter gene activity was significantly increased up to 1.8-fold compared to that of wild-type DH5α/pKMD3 in the exponential growth phase and reached its highest activity of 4.0 × 105 RLU/OD600 after 5 h of growth. Moreover, the observed effects were reduced in the DH5α Δhfq/pKMD3 + pLH01 strain to a level similar to that of the wild type. TS18/08 Δhfq/pKMD3 deletion mutants showed behavior similar to that of the DH5α deletion mutants. The relative reporter gene activity was significantly increased up to 2-fold in the late growth phase, showing its highest activity of 5.6 × 105 RLU/OD600 after 4 h of growth. The effects were reduced to the wild-type level after complementation of the TS18/08 Δhfq/pKMD3 strain with pLH01. Moreover, DH5α and TS18/08 did not show great differences in PsubAB1 activity in either complemented strains or empty vector controls (pBR322-harboring strains). This result indicated the pBR322 backbone had no effect on the PsubAB1 promoter activity.
FIG 4 Impact of Hfq on PsubAB1 activity in E. coli DH5α (A) and STEC E. coli TS18/08 (B). Relative reporter gene activity (RLU/OD600) is shown. Bars represent data from the wild-type strain (DH5α/pKMD3 [A] or TS18/08/pKMD3 [B], solid blue) and an empty vector control (DH5α/pKMD3 + pBR322 [A] or TS18/08/pKMD3 + pBR322 [B], dashed blue). Additionally, data from E. coli DH5α Δhfq/pKMD3 (DH5α Δhfq) (A) and E. coli TS18/08 Δhfq/pKMD3 (TS18/08 Δhfq) (B) (solid green) deletion mutants and DH5α Δhfq/pKMD3 + pLH01 (DH5α Δhfq + hfq) (A) and TS18/08 Δhfq/pKMD3 + pLH01 (TS18/08 Δhfq + hfq) (B) (dashed green) complemented strains are depicted. Data are presented for measurements over 5 h of cultivation in LB at 37°C with aeration. Data from three biological replicates are shown; error bars indicate standard deviations from mean values. Statistical significance compared to the wild-type strain is indicated (*, P < 0.05; **, P < 0.01).
The deletion of hfq led to a significant increase of PsubAB1 activity in the late growth phase for both E. coli DH5α and STEC TS18/08. Moreover, observed effects were restored to the wild-type level if complementation was performed. These results indicate that Hfq represses subAB1 promoter activity under wild-type conditions.

PsubAB1 activity is dependent on H-NS in E. coli strains.

The impact of deletion of the hns gene on PsubAB1 activity is depicted in Fig. 5. Relative reporter gene activity was significantly increased to a value of 1.9 × 105 RLU/OD600 in the DH5α Δhns/pKMD3 laboratory strain compared to that of the control DH5α/pKMD3 after 1 h of cultivation. After 2-, 3-, and 4-h time points, the PsubAB1 activity of the DH5α Δhns/pKMD3 strain was similar to that of the corresponding wild type. However, in the late exponential growth phase, the activity decreased up to a significant reduction of 1.5-fold relative reporter gene activity compared to that of the control. Nevertheless, the effects detected in the deletion mutant were restored in the complemented strain to levels comparable to that of E. coli DH5α.
FIG 5 Impact of H-NS on PsubAB1 activity in E. coli DH5α (A) and STEC TS18/08 (B). Relative (rel.) reporter gene activity (RLU/OD600) is shown. Bars represent data from the wild-type strain (DH5α/pKMD3 [A] or TS18/08/pKMD3 [B], blue) and an empty vector control (DH5α/pKMD3 + pBR322 [A] or TS18/08/pKMD3 + pBR322 [B], dashed blue). Additionally, data from E. coli DH5α Δhns/pKMD3 (DH5α Δhns) (A) and E. coli TS18/08 Δhns/pKMD3 (TS18/08 Δhns) (B) (solid red) deletion mutants and DH5α Δhns/pKMD3 + pLH01 (DH5α Δhns + hns) (A) and TS18/08 Δhns/pKMD3 + pLH01 (TS18/08 Δhns + hns) (B) (dashed red) complemented strains are depicted. Data are presented for measurements over 5 h of cultivation in LB at 37°C with aeration. Data from three biological replicates are shown; error bars indicate standard deviations from mean values. Statistical significance compared to the wild-type strain is indicated (**, P < 0.01).
The increase of PsubAB1 activity after 1 h of cultivation in the hns deletion mutant was not detected in STEC strain TS18/08, but, comparable to the case in E. coli DH5α, the deletion of hns led to a decrease of PsubAB1 activity. In the TS18/08 Δhns/pKMD3 strain, the decrease in relative reporter gene activity was measured during the whole period of cultivation, reaching a decrease of up to 2.9-fold compared to that of the wild-type strain. As described above for DH5α, the plasmid-based complementation led to a reduction of those effects.
These results suggested an impact of the global regulator H-NS on PsubAB1 activity. Repression of subAB1 by H-NS in early growth phase and activation in the exponential growth phase were measured for E. coli DH5α. In STEC strain TS18/08, only activation of the subAB1 promoter activity was detected.

The global regulators Hfq and H-NS control expression of subAB1, stx2a, and cdt-V.

Since TS18/08 was previously shown to express the three toxin genes subAB1, stx2a, and cdt-V, transcription analysis of wild-type and mutant strains was carried out by qRT-PCR. The gene expression of the three toxins was analyzed and normalized to expression of the reference gene rrsBCT value, where CT is threshold cycle). Transcription of all investigated strains was normalized to transcription of TS18/08 (ΔΔCT value) at each time point separately. The effect of the deletion of the hfq gene (Fig. 6A) and the hns gene (Fig. 6B) in E. coli TS18/08 is shown as fold change of mRNA levels in Fig. 6. The expression of subAB1 was increased significantly up to 2-fold in the E. coli TS18/08 Δhfq strain in the late exponential growth phase. The transcription of stx2a in this strain showed a similar pattern, whereas a 2.8-fold increase was measured after 5 h of cultivation compared to the level for TS18/08. The highest upregulation was detected for cdt-V expression. Similar to that of the other toxins, the 14-fold upregulation was measured in the late growth phase.
FIG 6 Expression of subAB1, stx2, and cdt-V in E. coli TS18/08 Δhfq (A) and TS18/08 Δhns (B) strains. Transcription is shown as fold change relative to the level of the control (E. coli TS18/08). Gene expression was measured under standard batch conditions (LB medium, 37°C, with aeration) using quantitative real-time PCR. Quantification was conducted using rrsB as a reference gene. Error bars indicate standard deviations from the mean values. Data from three biological replicates are shown. Asterisks indicate significance in fold change compared to the level for the control (P < 0.05).
Upregulation of the expression of all three toxins was also found in the E. coli TS18/08 Δhns strain. subAB1 transcription increased up to 6-fold after 1 h of cultivation and up to 3-fold in the late growth phase at 5 h of cultivation. The expression of cdt-V increased up to 16-fold in the late growth phase and therefore was comparable to the one obtained in the E. coli TS18/08 Δhfq strain. The expression of stx2a increased over the whole bacterial life cycle in the E. coli TS18/08 Δhns strain and was 10-fold upregulated after 2 h of cultivation.
The results obtained by qRT-PCR showed that both Hfq and H-NS have an impact on the expression of the toxins produced by STEC TS18/08. The strongest effects were detected in the late growth phase. In comparisons of the expression of the three toxins, the highest increase was detected in the E. coli TS18/08 Δhns for stx2a transcription. Therefore, the results of these experiments let us conclude that Hfq and H-NS had a strong impact on toxin gene transcription in STEC TS18/08.


Regulation of transcription of virulence factors is of major importance for pathogens, since maximum transcription of such factors at the site of infection may be advantageous for the colonization and infection process. In recent years, STEC strains have been analyzed frequently for the presence and absence of genes encoding virulence factors, such as Shiga toxins, type III effectors, adhesins, and EHEC-hemolysin, or for the qualitative description of their expression (14, 2730). In some cases, data on the regulation of virulence factors have been published. For example, Shiga toxin expression is included in the phage induction cycle and is recA dependent (31). Moreover, the complex regulation of the LEE operon is well elucidated, and there is some knowledge on regulation of EHEC-hemolysin.
During recent years, STEC strains that carry genes encoding a subtilase cytotoxin have been described increasingly, and it was demonstrated that this toxin contributes to virulence of STEC (11). In a recent study, we could show that the subtilase is expressed during the bacterial growth cycle in batch culture, but the dependence of regulators is currently not known (12). Therefore, we were interested in getting deeper insight into the transcription regulation of subAB1.
In this study, we investigated the effects of hfq and hns deletions in laboratory strain E. coli DH5α and wild-type STEC O113:H21 strain TS18/08. Both strains showed their highest promoter activity in the exponential growth phase. Hauser et al. (12) have demonstrated that transcription of subAB1 was highest after 3 h of cultivation under standard batch conditions, showing a 7-fold increase in transcription compared to the initial value. This is similar to the results obtained in our study. Interestingly, the promoter activity varied only slightly between the two tested strains.
Moreover, we could show that Hfq acts as a repressor of subAB1 promoter activity in the late growth phase. This finding was strain independent. Hfq is a pleiotropic regulator and has an impact on the expression of many virulence genes (23, 3235). In former studies, the regulation of toxin gene expression by Hfq was shown. Kendall et al. studied EHEC O157:H7 strain 86-24 and its mutant with a hfq deletion and its respective Shiga toxin expression. They could show that transcription of stx2AB was increased in this deletion mutant (23). This result could be reproduced in our study with the E. coli TS18/08 Δhfq strain. In addition, upregulation of cdt-V was detected in the same mutant strain, assuming that Hfq is integrated in the regulation of different toxins. Both SubAB and Shiga toxin belong to the family of AB5 toxins (36, 37), whereas the Cdt-V toxin is an AB2 toxin consisting of three subunits (Cdt-A, Cdt-B, and Cdt-C) (38). The regulation by Hfq seems not to be dependent on the type of toxin.
Hfq can directly interact with DNA and regulate transcription, but most processes described indicate regulation by Hfq-dependent small regulatory RNAs (sRNAs) (25, 34, 39). In recent studies, the prevalence of sRNAs in gene regulation were shown (40). Oogai et al. have shown that the regulation of expression of Cdt-V is dependent on sRNAs in Aggregatibacter actinomycetemcomitans (41). We suggest that this is the same regulatory pathway in STEC strain TS18/08. Regulation of toxins could be integrated in networks regulated by sRNAs due to the fast response to environmental changes. This cascade is controlled by the RNA chaperone Hfq.
Regulation of toxin gene expression was reported not only for sRNAs and Hfq but also for H-NS. Wan et al. revealed, by RNA sequencing, the repressing role of H-NS on expression of the hemolysin and Shiga toxin in EHEC (42). In our study, we showed by quantitative real-time PCR that H-NS repressed the expression of not only stx2a but also cdt-V and subAB1. Upregulation of subAB1 expression in the TS18/08 Δhns strain was detected in the early and late growth phases. For the E. coli DH5α Δhns strain, upregulation could be confirmed only in the early growth phase. Moreover, we measured a decreased promoter activity in the latter strain in the late growth phase. This result might be due to the different methods applied, and factors such as RNA decay could influence the results. Another possibility is that the toxin genes studied are not regulated directly by H-NS but are integrated in the regulatory pathway of H-NS. Wan et al. identified, by RNA sequencing, up to 983 genes which are regulated by H-NS in EHEC (42). H-NS is an important factor in xenogeneic gene silencing. Genes involved in response to environmental stimuli, such as the acidic environment, temperature, pH value, or host signal peptides, could have a direct impact on promoter activity and posttranscriptional processes. Moreover, H-NS interacts directly with other nucleoid binding proteins, such as the DNA binding protein StpA (24). Those proteins regulate gene expression if H-NS is not present (43, 44). Moreover, it has been shown that StpA in combination with Hfq could reduce effects of an hns deletion mutant. Thus, it was shown that phenotypic characteristics of a Δhns mutant were counteracted by hfq on a multicopy plasmid (45). This counteraction by Hfq and StpA might be the reason why different results were acquired for subAB1 expression in the TS18/08 Δhns strain in this study by comparing the luciferase reporter gene assay and qRT-PCR.
Given the fact that both global regulators showed influence on gene expression of three important STEC toxins, the role of these proteins in virulence and pathogenicity of STEC should be further investigated.


In this study, we demonstrated that expression of subAB1, stx2a, and cdt-V is dependent on functional global regulatory proteins Hfq and H-NS in both E. coli DH5α and STEC strain TS18/08. The presented data reveal that the expression of the virulence factors is integrated in the circuit of global regulatory proteins. Further studies will be necessary to elucidate the complex regulatory network.


Bacterial strains and plasmids.

Bacterial strains and plasmids used in this study are shown in Table 1. Strains were routinely grown in Luria-Bertani (LB) broth consisting of 1.0% (wt/vol) tryptone, 0.5% (wt/vol) yeast extract, and 1.0% (wt/vol) sodium chloride, adjusted to pH 7.0. For solid media, 1.5% (wt/vol) agar-agar was added. If necessary, ampicillin, kanamycin, and tetracycline were added to the medium in final concentrations of 100 μg/ml, 50 μg/ml, and 10 μg/ml, respectively. Overnight cultures were prepared by incubating two colonies in 10 ml LB medium in 100-ml Erlenmeyer flasks at 37°C on a rotary shaker at 180 rpm.
TABLE 1 Strains and plasmids used in this study
Strain or plasmidRelevant geno- and phenotypeaReference or source
E. coli  
    DH5αtonA lacZΔM15 endA1 recA1 thi-1 supE44 phoA gyrA96 hsdR17 Δ(lacZYA-argF)U169 relA1Invitrogen
    DH5α ΔhnstonA lacZΔM15 endA1 recA1 thi-1 supE44 phoA gyrA96 hsdR17 Δ(lacZYA-argF)U169 relA1 ΔhnsThis study
    DH5α ΔhfqtonA lacZΔM15 endA1 recA1 thi-1 supE44 phoA gyrA96 hsdR17 Δ(lacZYA-argF)U169 relA1 ΔhfqThis study
    HB101F mcrB mrr hsdS20(rB mB) recA13 leuB6 ara-14 proA2 lacY1 galK2 xyl-5 mtl-1 rpsL20 (Smr) supE44 λ52
    TS18/08O113:H21, cdt-V+ subAB1+ stx2+26
    TS18/08 ΔhnsO113:H21, cdt-V+ subAB1+ stx2+ ΔhnsThis study
    TS18/08 ΔhfqO113:H21, cdt-V+ subAB1+ stx2+ ΔhfqThis study
    pKD46Contains genes of red recombinase system under control of araB promoter, Ampr46
    pKD4Template plasmid for kanamycin resistance cassette, Ampr, Kanr46
    pCP20Contains genes for FLP recombinase, Ampr46
    pKMD3Luciferase reporter gene plasmid containing the subAB1 promoter regionThis study
    pBR322Cloning vector, pMB1 origin of replication, Ampr, Tetr53
    p3121Luciferase reporter gene (luc) template vector, Ampr, Kanr54
    pWSK29Cloning vector (low copy number), Ampr55
    pLH01Complementation plasmid, pBR322 backbone, with hfq gene under control of its own promoter, TetrThis study
    pLH02Complementation plasmid, pBR322 backbone, with hns gene under control of its own promoter, TetrThis study
Ampr, ampicillin resistance gene; Kanr, kanamycin resistance gene; Tetr, tetracycline resistance gene.
Plasmids were prepared using a QIAprep spin miniprep kit (Qiagen, Germany) by following the manufacturer’s recommendations (QIAprep miniprep handbook from July 2006; Qiagen). Genomic DNA for cloning procedures was isolated using a DNeasy Blood and Tissue kit (Qiagen) according to the procedure described in the DNeasy Blood and Tissue handbook (July 2006). Concentration and purity of nucleic acids were determined using a NanoDrop 2000 spectrophotometer (ThermoScientific, USA) and by agarose gel electrophoresis.

Construction of deletion mutants.

Deletion of genes was conducted as described by Datsenko and Wanner (46), with minor modifications. Briefly, the genes of interest were exchanged with a kanamycin resistance cassette by homologous recombination. Primers were designed with Serial Cloner, version 2.6.1 (SerialBasics; Franck Perez, Paris, France). Whole-genome sequences of E. coli DH5α (NCBI accession number NZ_JRYM01000001), E. coli K-12 MG1655 (GenBank accession number NC_000913.3), and E. coli TS18/08 (NCBI accession number ASM199093v1) were used to design primers targeting hfq and hns, including their surrounding regions. Moreover, the genomic backgrounds of hfq and hns were compared in both strains by DNA sequencing. Thus, surrounding regions, meaning 86-bp upstream and 76-bp downstream sequences for the gene hfq and 144-bp upstream and 300-bp downstream sequences for the gene hns, were amplified using the primers stated in Table 1. Substitution of the kanamycin cassette was carried out using the helper plasmid pKD46, encoding the recombinase genes γ, β, and exo. Using a second helper plasmid, pCP20, which encodes FLP recombinase, the resistance cassettes were excised. Transformation of plasmids and linear fragments was performed as described below. Primers used for mutagenesis are shown in Table 2. Deletion mutants were verified by PCR with subsequent agarose gel electrophoresis and DNA sequencing. The E. coli DH5α Δhns, E. coli DH5α Δhfq, E. coli TS18/08 Δhns, and E. coli TS18/08 Δhfq deletion mutants were analyzed for a presumable 2nd copy of these genes by PCR using primers hfq_for, hfq_rev, hns_for, and hns_rev (Table 2).
TABLE 2 Oligonucleotide primers used in this study
Primer and/or categoryUseNucleotide sequencea (5′ to 3′)Source
Hfq_forDetection of hfq geneATGGCTAAGGGGCAATCTTTACAAGThis study
Hfq_revDetection of hfq geneTTATTCGGTTCTTCGCTGTCCTGThis study
Hns_forDetection of hns geneTTATTGCTTGATCAGGAAATCGTCGThis study
Hns_revDetection of hns geneATGAGCGAAGCACTTAAAATTCTGThis study
    Del_hfq_forDeletion of hfq gene, creation of resistance cassetteAATGTGTACAATTGAGACGTATCGTGCGCATTTTTTCAGAATCGCGATTGTGTAGGCTGGAGCThis study
    Del_hfq_revDeletion of hfq gene, creation of resistance cassetteAGCGTATAACCCTCTAAATAGATCAGCGGGGAACGCAGGATCCATGGTCCATATGAATATCCTCCThis study
    Hfq + 186up_forVerification of deletion of hfqCCTGGCTGCGTGGTTGGGAAGThis study
    Hfq-176down_revVerification of deletion of hfqACCAGAGATTCAAACTCCTGGAGGTCThis study
    Del-hns_forDeletion of hns gene, creation of resistance cassetteACATCCGTATCGGTGTTACCACGAAACGGCGTTGAGCAATCGCGATTGTGTAGGCTGGAGCThis study
    Del-hns_revDeletion of hns gene, creation of resistance cassetteATAGGGAATTCTCGTAAACACAACTAATACAGAAGACTGAAAGGCATGGTCCATATGAATATCCTCCThis study
    Hns + 244up_forVerification of deletion of hnsTAGGTTACATGCAGGCCTTCGTTGThis study
    Hns-705down_revVerification of deletion of hnsACAGTGCGCATGCCGCGTTCCThis study
Construction of complementation plasmids   
    Hfq-76down_PvuI_revComplementation studiesGGGGGcgatcgAGCGTATAACCCTCTAAATAGATCAGCGGGGAACGCAGGATCThis study
    Hns + 144up_AseI_forComplementation studiesCCCCCCaattaatACATCCGTATCGGTGTTATCCACGAAACGGCGTTGAGCAATCThis study
    Hns-300down_PvuI_revComplementation studiesCCCCCCcgatcgATAGGGAATTCTCGTAAACACAACTAATACAGAAGACTGAAAGGThis study
Construction of luciferase reporter gene plasmid pKMD3   
    subAB-gibson_forCloning of subAB1 promoter region for pKMD3TTCGCTATTACGCCAGCTGAACGGTATCGATCCGThis study
    subAB-gibson_revCloning of subAB1 promoter region for pKMD3GTCTTCCATAAGCTCCTCCAGTTAGTACThis study
    luc-gibson_forCloning of luc for pKMD3AGGAGCTTATGGAAGACGCCAAAAACThis study
    luc-gibson_revCloning of luc for pKMD3CCGATTCATTAATGCAGTCACAATTTGGACTTTCCGThis study
    Gibson-1_forVerification of pKMD3ACACCCGCCGCGCTTAATGCThis study
    Seq 2-gibson_revVerification of pKMD3GATACCGCTCGCCGCAGCCGThis study
    Seq 3-gibson_forVerification of pKMD3GGTTGTGGATCTGGATGCCGThis study
    Seq 4-gibson_revVerification of pKMD3AGGGCGTATCTCTTCATAGCThis study
Underlined bases depict homologous regions in mutagenesis, and underlined lowercase letters depict restriction sites for cloning strategy.


Transformation of PCR products and plasmids was conducted by electroporation (GenePulser Xcell electroporation system; Bio-Rad, USA), and competent cells were prepared as described previously (47). To each cell aliquot, 300 ng of PCR products or 30 ng of plasmid DNA was added. Transformation was performed in electroporation cuvettes (2 mm; Bio-Rad) at 25 μF, 200 Ω, 2.5 kV, and 5 ± 0.2 ms. Residual steps were carried out as described previously (47).

Construction of reporter gene plasmid pKMD3.

The luciferase reporter gene plasmid pKMD3 was constructed using the isothermal Gibson Assembly (NEB, USA) strategy. Therefore, a 405-bp region upstream of subAB1, including the putative subAB1 promoter region, was cloned from E. coli O113:H21 strain TS18/08. The plasmid map is shown in Fig. 7. The vector backbone consisted of plasmid pWSK29, which was linearized by restriction with PvuII and subsequent purification by agarose gel extraction using a QIAeX II gel extraction kit (Qiagen, Germany) according to the manufacturer’s instructions (QIAeX II handbook, March 2015; Qiagen). The luciferase gene (luc) insert (1,653 bp) was amplified using plasmid p3121 as the target using primers luc-gibson_for and luc-gibson_rev (Table 2). The promoter region of the subAB1 gene was amplified with primers subAB-gibson_for and subAB-gibson_rev (Table 2) using genomic DNA of E. coli TS18/08. The isothermal assembly was performed at 60°C for 50 min with an insert-to-vector ratio of 2:1. The assembly mixture was directly transformed by electroporation in competent E. coli DH5α cells as described previously. Cloning of the reporter gene plasmid was verified by PCR and DNA sequencing.
FIG 7 Plasmid map of the luciferase reporter gene plasmid pKMD3. The putative promoter region of subAB1 (405 bp; PsubAB1) was cloned upstream of the luciferase reporter gene luc (1,652 bp) in the pWSK29 backbone restricted with PvuII (pWSK29_PvuII).

Luciferase assay.

Analysis of subtilase cytotoxin promoter activity (PsubAB1) was conducted using a luciferase reporter system (Promega, Madison, WI) by following the recommendations of the manufacturer (luciferase assay system, technical bulletin from 2015; Promega Corporation). Strains harboring the reporter gene plasmid pKMD3 (Table 1) were grown in LB medium at 37°C, 180 rpm, for 16 h. Twenty-four milliliters of LB medium in a 100-ml Erlenmeyer flask was inoculated with an overnight culture to an initial OD600 of 0.1 (unless stated otherwise) and further incubated at 37°C, 180 rpm, for 5 h. During cultivation, hourly sample taking and measurement of the OD600 was conducted in duplicates for each culture. In addition, samples were collected to measure luciferase activity. Thus, 400-μl samples were taken in duplicates for each strain and centrifuged at 6,000 × g, 4°C, for 5 min. Supernatants were discarded and pellets were resuspended in 90 μl of 1× cell culture lysis reagent (CCLR; Promega, USA) containing 10 mg/ml bovine serum albumin (albumin fraction V; Carl Roth, Germany). Subsequent adding of 90 μl 1× CCLR containing 10 mg/ml lysozyme (Sigma-Aldrich, Germany) was performed, and samples were frozen immediately at –70°C. Thawing of samples was conducted by incubation at 23°C with 300 rpm for 20 min in a heating block.
Measurement of luciferase activity was performed as described here and prepared in duplicates for each sample. Fifty-microliter aliquots of thawed samples were transferred to a white microtiter plate (LUMITRAC 600; Greiner). As a substrate, 50 μl of luciferase assay reagent (Promega) was added to samples, and luminescence was measured immediately at 562 nm using a microplate reader (Infinite M200; Tecan). To compare the results during cultivation, relative reporter gene activity was defined as the ratio of relative light units (RLU) to the respective OD600. All experiments were performed three times, each on a different day.

Cloning of vectors for complementation experiments.

For complementation experiments, plasmids were constructed containing hfq and hns, including their putative promoter regions, as shown in Fig. 8. The medium-copy-number plasmid pBR322 (Table 1) was chosen as a cloning vector due to its compatibility with the luciferase reporter gene plasmid pKMD3. The plasmid was double digested with restriction enzymes PvuI (NEB, USA) and AseI (NEB, USA). The reaction was carried out using 1 μg of DNA template, 10 U of each enzyme, and NEB buffer 3.1. Restriction was performed at 37°C for 60 min. Inserts of the sequences were cloned, 86-bp upstream and 76-bp downstream sequences for the gene hfq and 144-bp upstream and 300-bp downstream sequences for the gene hns, and were amplified using primers with restriction sites at their 3′ ends as shown in Table 2. Amplification was conducted using genomic DNA of E. coli strain DH5α (DNeasy Blood and Tissue kit; Qiagen, Germany) as the template. Oligonucleotides (Eurofins MWG Operon, Germany) were designed using Serial Cloner, version 2.6.1 (SerialBasics; Franck Perez, Paris, France). Purification of linearized vector DNA was conducted by excising the respective band on an agarose gel (1%, wt/vol) and subsequently isolated using a Wizard SV gel and PCR clean-up system (Promega, USA). The amplified inserts were digested with PvuI and AseI as stated above and purified using a PCR purification kit (Qiagen, Germany) by following the manufacturer’s recommendations (QIAquick spin handbook, April 2015; Qiagen). Ligation of vector and the respective insert was performed with T4 DNA ligase (ThermoFisher Scientific) at 22°C for 1 h using a vector-to-insert ratio of 1:5. After ligation, the resulting plasmids pLH01 and pLH02 were transformed in competent E. coli DH5α cells as described previously, and transformants were selected on agar plates containing tetracycline (10 μg/ml). Recombinant plasmids were then verified by DNA sequencing using the respective primers listed in Table 2 and finally transformed in reporter strains and deletion mutants using electroporation as described previously.
FIG 8 Cloning strategy of complementation vectors pLH01 (A) and pLH02 (B). Positions of hns and hfq genes refer to the genome sequence of E. coli DH5α (NCBI accession number NZ_JRYM01000001).

Analysis of gene transcription on complementation plasmids.

The expression of the cloned genes on the recombinant plasmids used for complementation was verified by transcription analysis as described earlier (48). Briefly, RNA was isolated using the RNeasy minikit (Qiagen, Germany). Strains harboring complementation plasmids were grown in LB medium at 37°C with agitation to an OD600 of 1.0. Five hundred microliters of the culture was added to 1 ml of RNAprotect bacterial reagent (Qiagen, Germany). Isolation of RNA was conducted by following the manufacturer’s recommendation, and DNA was digested using a DNA-free kit (Thermo Fisher Scientific, USA) according to the manual (user guide, October 2012; Life Technologies Corporation). RNA concentration and purity were determined spectrophotometrically using a NanoDrop 2000 device (Thermo Scientific, USA), followed by visual inspection on denaturing agarose gel electrophoresis (data not shown). Transcription to cDNA was performed using 1 μg RNA, a SuperScript II reverse transcriptase kit (Bio-Rad, USA), and primer for detection of the hfq (hfq_for and hfq_rev) or hns (hns_for and hns_rev) gene (Table 2) by following the manufacturer’s recommendations (manual of SuperScript II from 2010; Life Technologies Corporation). As a control, cDNA synthesis was performed using random primers as given in the SuperScript II reverse transcriptase kit (random hexamer oligonucleotides). For each approach, the reverse transcriptase negative control was applied to avoid false-positive results due to residual DNA present in samples. Synthesized cDNA was analyzed by PCR using primers specific for genes hfq and hns as stated above. Subsequently, amplification was investigated by agarose gel electrophoresis. Expression control was performed in three biological replicates independently, each on different days.

RNA isolation.

Erlenmeyer flasks containing 24 ml LB medium were inoculated with overnight culture to an OD600 of 0.1. E. coli TS18/08 was cultivated at 37°C, 180 rpm, for 5 h, and samples were taken hourly for RNA isolation. Therefore, 1.0 × 109 cells were transferred to a 2-fold volume of RNAprotect bacterial reagent (Qiagen, Germany). The suspensions were mixed immediately for 5 s, followed by an incubation at room temperature for 5 min. Centrifugation was then performed at room temperature at 5,000 × g for 10 min, and the supernatants were discarded. Pellets were stored at –70°C until further processing. RNA isolation was performed as described in the manual of the RNAprotect bacterial reagent (Qiagen, Germany) and RNeasy minikit (Qiagen, Germany) by following the protocol for enzymatic lysis and proteinase K digestion (protocol 4) and subsequent purification of total RNA (protocol 7). RNA purity and concentration were determined using a spectrophotometer (NanoDrop 2000; Thermo Scientific, USA), and integrity of RNA was examined using formaldehyde-based denaturing agarose gel electrophoresis.

Transcription analysis.

Prior to cDNA synthesis, residual DNA was degraded in the samples using a DNA-free kit (ThermoScientific, USA) by following the recommendations stated in the manual (user guide, October 2012; Life Technologies Corporation). Reverse transcription was performed using an iScript cDNA synthesis kit (Bio-Rad, USA) with the following procedure: 500 ng of the RNA samples was used for reverse transcription and incubated according to the protocol stated by the manufacturer (manual from 2000; Bio-Rad Laboratories). For each sample, a reverse transcription-negative approach was conducted as a control to detect residual DNA. Synthesized cDNA was used for quantitative real-time PCR in a CFX96 system (Bio-Rad, USA) by following the experimental procedure described previously (12). Therefore, 10 μl of SsoAdvanced universal SYBR green supermix (Bio-Rad, USA), 2.8 ng of cDNA, 0.75 μl of each primer (10 pmol/μl), and nuclease-free water to the end volume of 20 μl were used for each qRT-PCR approach. Primers and respective PCR conditions are shown in Table 3. Analysis was performed in duplicates and with three biological replicates. Analysis of data was performed using the 2−ΔΔCT method described by Pfaffl et al. (49, 50). Thus, expression data for genes subAB1, stx2, and cdt-V were normalized to expression of reference gene rrsB (16S rRNA gene) (ΔCT). Subsequently, ΔCT values were normalized to the values of E. coli TS18/08 (ΔΔCT). The data were analyzed for each point of time independently. For all targets, standard curves of recombinant plasmids were used to identify target efficiency as described previously (12, 51). No-template controls and reverse transcription-negative controls were applied in each approach and showed CT values after 35 to 40 cycles of PCR (data not shown). Each sample was analyzed in at least two technical replicates, and experiments were conducted in three independent experiments on different days (biological replicates).
TABLE 3 Oligonucleotide primers used for transcription analysis by qRT-PCR
PrimerNucleotide sequence (5’ to 3’ direction)PCR product size and conditionsReference
rrsB-forGCATAACGTCGCAAGACCAAA91 bp; 95°C, 10 s; 60.0°C, 30 s; 72°C, 10 s51
RTsubAB-2-forGCAGGTATTATGGGATGTCT177 bp; 95°C, 15 s; 60.3°C, 20 s; 72°C, 10 s12
Upper-stx2TCATATCTGGCGTTAATGGAGTTC99 bp; 95°C, 10 s; 52.4°C, 30 s; 72°C, 10 s51
RTCDTA-forCCGATGGTAACACGCAATG184 bp; 95°C, 10 s; 53.0°C, 30 s; 72°C, 10 s12

Statistical analysis.

Means from biological replicates were used for statistical analysis. Data were analyzed on normal distribution using the Grubb’s test. If data sets of two different time points measured were compared for one strain, paired Student's t test was used, and if data were not normally distributed, Mann-Whitney test was applied. Data sets of different strains at identical time points measured were analyzed by application of analysis of variance (ANOVA). The variance homogeneity was identified by residual plots (Q-Q-plots). If requirements were given, one-way ANOVA was performed using a post hoc Tukey test (α = 0.05). If requirements were not met, data were analyzed using Welch’s ANOVA and post hoc Games-Howell test. Statistical analysis was performed using the software SPSS Statistics 25 (IBM, USA), and statistical significance was given if the P value was <α.


We thank Lydia Pertschy and Markus Kranz (University of Hohenheim, Stuttgart) for skillful technical assistance. We gratefully thank Hans-Peter Piepho (University of Hohenheim, Stuttgart) for his advice on the statistical analysis of the data.
This work was funded by the German Research Foundation (DFG) grant number SCHM-1360/11-1.
We have no conflicts of interest to declare.

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Information & Contributors


Published In

cover image Applied and Environmental Microbiology
Applied and Environmental Microbiology
Volume 85Number 2015 October 2019
eLocator: e01281-19
Editor: Maia Kivisaar, University of Tartu
PubMed: 31375495


Received: 6 June 2019
Accepted: 30 July 2019
Published online: 1 October 2019


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  1. STEC
  2. Shiga toxin-producing Escherichia coli
  3. gene expression
  4. global regulator
  5. hfq
  6. hns
  7. subtilase cytotoxin



Laura Heinisch
Institute of Food Science and Biotechnology, Department of Food Microbiology and Hygiene, University of Hohenheim, Stuttgart, Germany
Katharina Zoric
Institute of Food Science and Biotechnology, Department of Food Microbiology and Hygiene, University of Hohenheim, Stuttgart, Germany
Maike Krause
Institute of Food Science and Biotechnology, Department of Food Microbiology and Hygiene, University of Hohenheim, Stuttgart, Germany
Herbert Schmidt
Institute of Food Science and Biotechnology, Department of Food Microbiology and Hygiene, University of Hohenheim, Stuttgart, Germany


Maia Kivisaar
University of Tartu


Address correspondence to Herbert Schmidt, [email protected].

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