Transcriptional cross-activation between toxin-antitoxin systems of Escherichia coli
© Kasari et al.; licensee BioMed Central Ltd. 2013
Received: 13 December 2012
Accepted: 18 February 2013
Published: 21 February 2013
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© Kasari et al.; licensee BioMed Central Ltd. 2013
Received: 13 December 2012
Accepted: 18 February 2013
Published: 21 February 2013
Bacterial toxin-antitoxin (TA) systems are formed by potent regulatory or suicide factors (toxins) and their short-lived inhibitors (antitoxins). Antitoxins are DNA-binding proteins and auto-repress transcription of TA operons. Transcription of multiple TA operons is activated in temporarily non-growing persister cells that can resist killing by antibiotics. Consequently, the antitoxin levels of persisters must have been dropped and toxins are released of inhibition.
Here, we describe transcriptional cross-activation between different TA systems of Escherichia coli. We find that the chromosomal relBEF operon is activated in response to production of the toxins MazF, MqsR, HicA, and HipA. Expression of the RelE toxin in turn induces transcription of several TA operons. We show that induction of mazEF during amino acid starvation depends on relBE and does not occur in a relBEF deletion mutant. Induction of TA operons has been previously shown to depend on Lon protease which is activated by polyphospate accumulation. We show that transcriptional cross-activation occurs also in strains deficient for Lon, ClpP, and HslV proteases and polyphosphate kinase. Furthermore, we find that toxins cleave the TA mRNA in vivo, which is followed by degradation of the antitoxin-encoding fragments and selective accumulation of the toxin-encoding regions. We show that these accumulating fragments can be translated to produce more toxin.
Transcriptional activation followed by cleavage of the mRNA and disproportionate production of the toxin constitutes a possible positive feedback loop, which can fire other TA systems and cause bistable growth heterogeneity. Cross-interacting TA systems have a potential to form a complex network of mutually activating regulators in bacteria.
Bacterial toxin-antitoxin (TA) systems are complexes of a stable toxic- or growth-arresting factor and its unstable inhibitor [1, 2]. They are diverse, abundant in all bacteria, except a few intracellular parasites, and are found in many archaea [3–6]. On the basis of their ubiquity and diversity, we can assume that regulation by TA must be common and beneficial in a wide range of microorganisms. However, their role in bacterial physiology is unclear [7, 8], in part due to redundancy . They were first discovered in plasmids and characterized as addiction systems, which are responsible for post-segregational killing . However, because of its high cost to the host, such a stability mechanism is used only in rare cases . Chromosomal TA loci were found thanks to full genome sequencing  and were demonstrated to be functional, expressed at significant levels, and activated by various stressful conditions, particularly by amino acid starvation [12–15].
Our current study focuses on type II TA systems. In this group, both the toxin and the antitoxin are proteins, which are encoded by adjacent co-transcribed genes. In a growing cell, toxins are neutralized by tightly bound antitoxins. Antitoxins are degraded by proteases much more quickly than toxins, and if antitoxin production stops, toxins target vital functions of the producer through diverse mechanisms. Many toxins (e. g. RelE, MazF, YafQ, HigB, HicA, MqsR) are endoribonucleases and inhibit protein synthesis through cleavage of free or ribosome-bound mRNA [16–21]. MazF also cleaves 16S rRNA  and VapC endonucleases of enteric bacteria cleave initiator tRNA . Another group of toxins (CcdB, ParE) interferes with DNA gyrase [24, 25], whereas HipA is a protein kinase [26, 27], and zeta toxins (PezT) inhibit cell wall synthesis . Activation of toxins causes growth inhibition and dormancy that may be transient  but in some circumstances is irreversible and leads to cell death [28, 30–32].
Besides direct protein-protein interaction, antitoxins regulate toxin activity at the level of transcription. Antitoxins are DNA-binding proteins and specifically repress transcription of their own TA operons both alone and, even more effectively, in complexes with their cognate toxins. Degradation of an antitoxin causes de-repression of the TA promoter  and allows the toxin activity to be detected indirectly by measurement of transcript levels. Gerdes and colleagues have demonstrated fine-tuning of transcription by the toxin:antitoxin ratio for the RelBE system [34, 35]. The RelB antitoxin in excess of the RelE toxin promotes formation of the RelB:RelE (2:1) complexes that bind to the operator sites and repress transcription. RelE toxin in excess promotes formation of the ReB:RelE (2:2) complexes that are unable to bind DNA . As a result, over-expression of RelE causes substantial increase in the relBE mRNA level. These authors suggested that such transcriptional regulation by the T:A ratio is commonplace for TA loci  and demonstrated it recently for VapBC . Importantly, the levels of TA mRNAs were increased in cell populations enriched for persisters, thereby linking TA systems to antibiotic susceptibility [38, 39]. Persisters are transiently dormant bacteria that remain non-dividing under growth-supporting conditions and are not killed by bactericidal antibiotics . TA systems, by their very nature, may be primarily responsible for persister formation. Mutations that increase toxicity of the TA toxins were shown to increase the frequency of persisters and cause high persistence phenotypes [41, 42]; and deletion of the yafQ toxin significantly decreased persister frequency in E. coli biofilms . A recent study reports that successive deletion of 10 endoribonuclease-encoding TA loci progressively reduced the level of persisters while single deletions of TA systems had no effect on persister frequency in planktonic E. coli. Hence, it is extremely important to consider redundancy and possible cross-talk when we study TA-related phenotypes, because most bacterial genomes contain multiple TA loci.
In the current study we found that uninhibited toxins can activate transcription of the other TA operons. Cleavage of these transcripts by endoribonuclease toxins adds another layer of complexity. Reciprocal transcriptional de-repression and transcript cleavage predict that toxin-antitoxin systems have a potential to form a complex network of regulators that controls growth and dormancy of bacteria.
As shown in Figure 1, we indeed saw a clear cross-activation of relBEF in response to all toxins tested except YafQ. Induction of RelE, MazF, MqsR, HicA and HipA conferred a clear increase in the relBEF mRNA level in an hour. Use of three separate probes revealed, however, that different mRNA species pile up in response to different toxins. Before induction and 15 min after, all three probes – relB, relE and relF – detected a transcript of the same size corresponding to the full-length mRNA of the operon , as confirmed later by primer extension mapping of the 5′ end (Additional file 1: Figure S4). Only after MazF expression a shorter transcript, a putative cleavage product, could be detected at the 15 min time point using relE probe (Figure 1B). At later time points, hybridization with relB (Figure 1A) and relE (Figure 1B) probes gave different signals: in response to induction of MazF, MqsR, and HicA we saw cleavage of the full-length mRNA and massive accumulation of the toxin-encoding part, while the antitoxin-coding portion could not be detected and was apparently degraded (Figure 1A,B). Such cleavage and accumulation of the toxin portion also occurred in response to RelE. Hybridization with relF probe revealed additional cleavage, both within relE and downstream, in response to expression of all these toxins, and the relF part accumulated as the most abundant portion of the relBEF transcript (Figure 1C). Also, some transcripts larger than the full relBEF mRNA appeared, particularly after induction of RelE and MqsR. Production of HipA, which is not a ribonuclease, conferred strong induction of full-length relBEF mRNA but cleavage and uneven accumulation of different mRNA fragments could not be seen. MUP treatment produced overproduction of the full relBEF mRNA as well as accumulation of some cleavage products. Production of YafQ did not lead to a clear cross-activation of relBEF transcription. However, relE probe showed accumulation of a short RNA fragment in response to this toxin. It is possible, that transcription of the operon is activated by YafQ but the transcript is degraded to small fragments. Clearly, these fragments cannot serve as templates for synthesis of RelE and, therefore, functional cross-activation does not occur. Modest induction of relBEF with no cleavage was evident in the 1h and 2h samples of control cultures, lacking artificial production of any free toxin. We have to consider that, at this stage, the control cultures were approaching stationary phase, and induction of toxin-antitoxin modules has been described in similar conditions .
Probes complementary to yiaF and rpsS were used for control because the levels of transcription of these genes did not differ between log phase cells and the ampicillin-refractory non-growing subpopulation, where TA operons were highly expressed . rpsS is a part of the large S10 ribosomal protein operon with an estimated transcribed length of 5181 bp ; yiaF (711 bp ORF) encodes for a putative membrane protein of unknown function; it is located between genes pointing in the opposite direction and must form a single-gene operon. The control mRNAs were not induced by toxins (Additional file 1: Figure S2B,C). After induction of toxins, the yiaF transcript was degraded without accumulation of any stable fragments. (Additional file 1: Figure S2B). Surprisingly, mupirocin initially induced transcription of yiaF whereas the level of the transcript dropped after longer incubation (Additional file 1: Figure S2B). The S10 transcript was degraded as well. Some accumulating stable fragments of the S10 transcript were detectable after MazF, RelE and MqsR production (Additional file 1: Figure S2C).
To be sure that the accumulating RNA fragments, which correspond to the 3′ portion of the relBEF mRNA, are not initiated from toxin-inducible cryptic promoters within the operon, we deleted the promoter of the relBEF operon. In the promoterless BW25113 ΔP relBEF strain, we did not see induction of the relBEF mRNA nor the characteristic accumulation of its 3′ portion (Additional file 1: Figure S3). We still saw a transcript that could be detected by the relE and relF probes (Additional file 1: Figure S3B,C) but the level of this transcript did not depend on the RelE production. It might be initiated from a constitutive promoter that was newly created by deletion of P relBEF . Transiently induced smear of RNA that was detected in BW25113 ΔP relBEF with the relB probe (Additional file 1: Figure S3A, lanes 6 and 7) is transcribed from the RelB-expression plasmid pKP3033. That is the reason why we omitted this plasmid when we studied induction of relBEF in response to RelE (Figure 1, Additional file 1: Figure S3, lanes 8–11). Thus, we can be sure that the shorter transcripts that massively pile up in response to toxins are indeed cleavage products and are initiated at the genuine P relBEF promoter.
Next, we tested whether over-production of the toxin RelE activates other toxin-antitoxin genes in the chromosome. The northern hybridization results show strong induction of the mqsRA, mazEF, dinJ-yafQ, hicAB, yefM-yoeB, and prlF-yhaV TA systems (Figure 2). Similarly to relBEF, the induced transcripts were cleaved and the toxin-encoding parts seem to accumulate preferentially while the antitoxin-coding parts are more effectively degraded. That appears to be true irrespective of whether the toxin is encoded by the first (mqsRA, hicAB) or the second (mazEF, yefM-yoeB, prlF-yhaV) gene of the operon (Figure 2). Reliable testing of this phenomenon requires characterization of the cleavage products and additional experiments in the future.
Additional experiments indicated that transcriptional cross-activation of TA operons does not occur between all possible TA combinations. Northern hybridization using mqsR probe showed that overproduction of MazF and HicA does not induce the mqsRA promoter while YafQ and HipA induce it (data not shown), as well as RelE (Figure 2).
To confirm our notion of TA cross-activation, we hoped to see some cleavage hotspots. At those sites, strong cleavage by an overproduced toxin occurs at its specific cutting sequence (e.g. ACA in the case of MazF). Cleavage at the same site in response to expression of another toxin would indicate activation of the primary cutter by the over-produced toxin. We tested possible cross-activation at three of these sites. At position 174 (ˇACA), the relBEF transcript is cut by MazF and in response to the over-produced HicA. The MqsR-specific cleavage sites at positions 399 (GCˇU) and 431 (GˇCU) are also cleaved in the samples from HicA over-production (Additional file 1: Figure S4). We found that these cuts were not due to the activation of MazF and MqsR, since they occurred in RNA extracted from the BW25113ΔmazEF and BW25113ΔmqsRA cells (data not shown). ChpBK, a homolog of MazF with similar but relaxed sequence specificity  may be accountable for the cleavage at 174 (ˇACA).
The toxin-encoding parts of the TA transcripts seem to be generally more stable than the antitoxin-encoding parts and accumulate after cleavage (Figures 1, 2). If the toxin open reading frame (ORF) on these cleavage products is intact and translated into a functional protein, the T:A balance must be shifted towards toxin followed by more cleavage, cross-activation of other TA systems, and inhibition of protein synthesis. That creates the possibility of a positive feedback circuit and even a network of them. A positive autoregulatory loop, in turn, could explain the bistability of bacterial growth observed in response to toxin expression [53, 54].
Production of toxins causes an extensive rearrangement of bacterial physiology. It can inflict dormancy and antibiotic tolerance  if the toxin level exceeds a threshold . Fluctuations in toxin levels above and below the threshold have been used to explain the coexistence of dormant and growing cells in a population . The possibility of positive feedback by the generation and selective buildup of the toxin-encoding mRNA fragments may explain this heterogeneity in growth. Therefore, we wanted to evaluate the recovery of single bacteria and test possible growth heterogeneity after over-production of a toxin and the resulting activation of the chromosomal TA loci. We monitored growth resumption by individual cells using dilution of previously synthesized green fluorescent protein (GFP) . The plasmid pTM11 was inserted into the chromosome of BW25311 to allow IPTG-inducible GFP to be expressed, and this strain was transformed with plasmids for L-arabinose-inducible production of toxins RelE, MazF, MqsR and HipA. Expression of GFP was induced for 2.5 h; thereafter, the cells were transferred into medium containing L-arabinose to induce the toxins. After 90 min, the growth medium was changed again to shut down toxin synthesis and allow recovery (Additional file 1: Figure S5). Analysis of the bacterial GFP content by flow cytometry (Additional file 1: Figure S6) showed that after temporary expression of RelE and HipA the bacteria resumed growth rather uniformly, while after expression of MazF and MqsR a subpopulation started to grow with a delay. Thus, expression of these toxins created bistability in a population. Most importantly, all bacteria resumed growth after the transient expression of toxins. Although inhibition by MazF and MqsR was apparently stronger and induced growth heterogeneity, it did not generate a subpopulation of persistently non-dividing bacteria (Additional file 1: Figure S6).
Induction of the chromosomal relBEF in response to the ectopically produced RelE can be explained by conditional cooperativity (dependence of transcriptional regulation on the T:A ratio) . However, according to our current knowledge, such mechanism is not applicable to cross-induction. Activation of YoeB by VapC depended on Lon protease . Also, Lon was required for induction of TA operons in response to amino acid starvation and chloramphenicol [14, 17, 18, 61]. Our experiments do not provide a solid support for the role of Lon and ClpP in cross-regulation between TA systems of E. coli (Figure 4). Since the cross-induction was present in the knock-out strains, an additional, Lon-, ClpP-, HslV-, and polyphosphate-independent mechanism of regulation must be involved. Unlocking this mechanism remains a task for future studies. The simplest explanation to activation of TA systems would be depletion of antitoxins. It must inevitably happen when protein synthesis decreases. That predicts nonselective induction of all TA operons in response to inhibition of translation, no matter if it is caused by starvation or artificial production of a toxin. Requirement of relBE for transcriptional activation of mazEF during amino acid starvation (Figure 3) contradicts this prediction as well as the lack of mqsRA induction in response to overproduction of MazF and HicA (data not shown). An option for a mechanism of cross-activation is positive feedback regulation due to selective accumulation of toxin-encoding fragments upon mRNA cleavage. As we saw, after cleavage by overproduced toxin, the antitoxin-encoding RNA fragments are rapidly degraded while the toxin-encoding fragments may serve as templates for translation of toxin. Different toxins produce different cleavage products. That can potentially explain why they cause unequal level of trans-activation when overproduced.
Another intriguing issue of TA cross-reaction is the possible cross-inhibition due to non-cognate interactions. Some authors report such cross-reactions [63–68] while others have tested but not found them [69, 70]. As a part of this study, we examined non-cognate inhibition between E. coli toxins and antitoxins of the RelBE, MazEF, MqsRA, and HipBA systems in vivo. In this attempt, we run into a previously described phenomenon that may become a source of erroneous results. If toxins are expressed from the arabinose-inducible P BAD promoter and antitoxins from an IPTG-inducible promoter, it is important to consider that IPTG inhibits P BAD directly . When we used an expression vector that encoded for the IPTG-insensitive C280* version of AraC transcriptional activator, we could not see any cross-inhibition. Based on that, a recent report on functional non-cognate TA interactions in Mycobacterium tuberculosis may require retesting.
In the current study, we found that the cleavage products produced by TA toxins differ in stability. Selective targeting of mRNAs by endoribonucleolytic toxins and different stabilities of the resulting cleavage products may constitute another layer of gene regulation in the bacterial stress response. Differences in half-life and translational efficiency of mRNA cleavage products, along with generation of a pool of ribosomes lacking the anti-Shine-Dalgarno sequence (as shown for MazF ), could profoundly affect the proteome composition. An example of such an effect is the occurrence of a MazF-resistant protein pool in E. coli. The accumulation of toxin-encoding mRNA fragments may have potential use as a marker of toxin activation in studies of stressed and non-growing bacteria. Increase of the T/A ratio may possibly trigger a positive feedback loop consisting of transcriptional activation of the TA operon, successive cleavage of the TA transcript, buildup of the toxin-encoding mRNA fragments, and translation of them, shifting the T/A balance (Figure 7). Thus, it can be related to TA-linked growth heterogeneity in bacterial populations (Additional file 1: Figure S6) [38, 39, 54].
The main finding of this study is that bacterial toxin-antitoxin systems affect mutually each others’ expression and activity (Figure 7). We show that overexpression of one toxin can activate transcription of the other TA operons. Toxins with endoribonuclease activity add another layer of complexity to these interactions. They cleave TA mRNA, which is followed by degradation of the antitoxin-encoding RNA fragments and accumulation of the toxin-encoding fragments. We show that these accumulating mRNA fragments can be translated to produce more toxin.
Most of bacteria have many different TA systems. Although their function is debatable, many TA toxins have similar activity and the inhibitory effect on bacterial cells is common to all of them. Therefore, an important question is whether TA systems are redundant or not. Another intriguing issue is whether different TA systems are functionally connected and do cross-talk [44, 70].
Here we over-expressed toxins to show that TA systems have a potential to form a network of cross-reacting regulators in E. coli. We found an example of such cross-reaction, which occurs without artificial overexpression: the relBE-dependent transcriptional activation of mazEF during amino acid starvation. It remains a rather difficult task to identify the mechanism(s) of TA cross-activation. Currently we know that cross-activation is not dependent on major proteases Lon, ClpP, and HslV. Also, it cannot be a self-evident outcome of antitoxin shortage since we know examples where shutdown of protein synthesis does not activate a TA promoter.
All strains and plasmids are listed in Additional file 1: Table S1. Conditions of bacterial cultivation and construction of strains and plasmids are described in Additional file 1: Supporting information.
Procedures for blotting and hybridization are described in . E. coli BW25113 was transformed with two plasmids, one bearing an antitoxin gene and the other bearing a toxin gene. Cultures containing the empty vector plasmids pBAD33 and pOU82 were used for negative controls. When bacteria contained plasmids for toxin expression, the LB medium for overnight cultures was supplemented with 0.2% glucose and 50 μM IPTG (for HicA with 1mM L-arabinose). Overnight cultures were diluted 1000-fold into 200 ml of LB and grown to OD600 ≈ 0.2 (for ~ 2.5 h). To induce toxins, 1 mM L-arabinose, 1 mM IPTG (for HicA) or 30 μg ml−1 mupirocin was added. Overnight cultures of BW25113 ΔrelBEF and BW25113 ΔP relBEF containing plasmids were diluted into LB supplemented with 0.2% glucose and 50 μM IPTG; at OD600 ≈ 0.2, bacteria were collected by centrifugation (5 min, 5000g, at 20°C) and resuspended in prewarmed LB supplemented with 1 mM L-arabinose. Total RNA was extracted using two different protocols: in Figures 2, 6 and S3 we used Trizol reagent  and in all other experiments we used hot phenol (for details see Additional file 1: Supporting information). Samples of total RNA (10 μg) were subjected to electrophoresis on denaturing gels. The DNA oligoprobes used for hybridization are listed in Table S2 (Additional file 1). For re-hybridization, the membranes were stripped by boiling for 2×10 min in 0.1% SDS, 5mM EDTA. Chemiluminescent signals were captured using ImageQuant RT ECL imager (GE Healthcare) and X-ray film (Agfa).
RNA samples were collected as for northern blotting. Extension primers (Additional file 1: Table S2) were labeled with [γ32P]ATP by T4 polynucleotide kinase (Thermo Scientific) and purified with a Nucleotide Removal Kit (Qiagen). Total RNA (15 μg) was mixed with labeled primer and incubated at 75°C for 2 min followed by slow cooling for 25 min. Extension reactions were carried out at 44°C for 30 min using 200U of RevertAidTM H minus reverse transcriptase (Thermo Scientific) and stopped with 10 μl of formamide loading buffer . Reaction products were concentrated by ethanol precipitation before gel electrophoresis. DNA was sequenced using a Sequenase Version 2.0 Kit (USB Products, Affymetrics). A PCR product amplified using primers relBEFup and relFdwn, and treated with Exonuclease I and shrimp alkaline phosphatase (ThermoScientific), was used as the template for the sequencing reactions. Samples were analyzed by 7M urea-6% polyacrylamide gel electrophoresis.
To prepare lysates, bacteria were grown to an OD600 of ~0.7 and expression of T7 RNA polymerase was induced for 1 h by adding 1mM IPTG. Control cultures were grown without IPTG. Bacteria were spinned down and lysed in Laemmli sample buffer. Proteins were separated by tricin–SDS–13% polyacrylamide gel electrophoresis . For detection of the His6-tagged toxins, the proteins were electroblotted onto Hybond-ECL nitrocellulose membrane filters (GE Healthcare) and probed with nickel-activated horseradish peroxidase (HisProbeTM-HRP; Thermo Scientific).
Overnight cultures were grown from fresh single colonies for 17–18 h in LB supplemented with 0.2% glucose and diluted 500-fold, into 3 ml of broth. After 2 h of incubation, 1 mM IPTG was added to initiate synthesis of green fluorescent protein (GFP). Expression of GFP was induced for 2.5 h. Then, cells were collected by centrifugation and transferred into LB supplemented with 0.2% L-arabinose to induce toxin synthesis. During the change of the medium, the culture was diluted 10-fold. After 90 min, the growth medium was changed to LB containing 0.2% glucose to stop the production of toxins, this time with 2-fold dilution. Starting from the induction of toxin synthesis, samples were taken for flow cytometry analysis and OD600 measurement. For flow cytometry analysis, the samples were mixed with an equal volume of 30% glycerol in PBS and stored at −80°C pending analysis. After dilution with sterile PBS, the samples were analyzed using an LSRII and a high-throughput sampler (BD) with a laser beam maximum wavelength of 488 nm. The results were analyzed by using FlowJo 7.2.1software.
All growth inhibition (Additional file 1: Figure S1) and growth resumption experiments (Additional file 1: Figure S5, S6) were repeated at least three times. All northern blot (Figures 1, 2, 3, 4, 6 Additional file 1: Figures S2, S3), primer extension mapping (Additional file 1: Figure S4) and in vivo translation experiments (Figure 6) were repeated at least twice with similar results. Typical results are presented for these experiments and for the FACS analysis of growth resumption (Additional file 1: Figure S6).
This work was supported by Estonian Science Foundation grant 8822 and by the European Regional Development Fund through the Center of Excellence in Chemical Biology. We thank Kenn Gerdes, Edita Sužiedėlienė, and Kim Lewis for plasmids and strains; and Vasili Hauryliuk, Ülo Maiväli, Isabella Moll and Arvi Jõers for comments on the manuscript. The authors would like to thank the anonymous reviewer who suggested the mupirocin experiment as a test for the TA cross-activation in physiologically relevant conditions.
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