Identification of GcvB as a Regulator ofphoP Expression

Even though MicA is an Hfq-dependent sRNA, expression ofphoP was found to be strongly activated at the post-transcriptional level by the deletion ofhfq in both wt andmicA deleted cells[24]. We thus hypothesized that one or several Hfq-dependent regulator(s) could affectphoP expression independently of MicA. Therefore, we transformed a strain carrying a P_BAD-phoP-lacZ reporter fusion with a plasmid library overexpressing most of the knownE. coli Hfq-dependent sRNAs from an IPTG-inducible modified P_lac promoter[8]. Transcription of thephoP-lacZ fusion is driven by the P_BAD promoter so that expression of this fusion should not be sensitive to control ofphoP at the transcription initiation level. The transcription start site is expected to be identical to that of the proximalphoP promoter, P₁, which is normally positively regulated by PhoP inE. coli[26]. The fusion encompasses only 66 nts ofphoP mRNA, that correspond to a 36 nts 5′ leader followed by the first 30 nts of the ORF. The ß-galactosidase activity of the different transformants was assayed and the results are shown inFigure 1A. Of the 25 sRNAs tested, 4 modulated the expression of the fusion by more than 2-fold, SgrS and RydC positively and MicA and GcvB negatively. Since SgrS is involved in sugar metabolism[27], we suspected that its overproduction could affect expression from the arabinose-induced P_BAD promoter. To test this possibility, we measured the SgrS-mediated repression of the samephoP-lacZ fusion when constitutively expressed from the P_tet promoter instead of P_BAD. Since the pSgrS plasmid had no effect on this P_tet-phoP-lacZ fusion (Figure 1B), it is likely that it activated the P_BAD- driven fusion at the promoter level and this was not investigated further. The RydC sRNA has been shown to activate (repress) the expression of fusions that are negatively (positively) regulated by Hfq, most likely by titrating Hfq[28]. One possibility is therefore that it acts on P_BAD-phoP-lacZ in the same way, but further experiments are required for a definitive proof. The same may be true for sRNAs such as ChiX, that also activates the expression of the fusion almost 2-fold.

In the experiment shown inFigure 1A, pMicA repressed the expression ofphoP-lacZ by 3.1-fold, which is in agreement with our previous results. Furthermore, this experiment also identified GcvB as a multicopy repressor ofphoP-lacZ, since pGcvB was responsible for a 4.5-fold decrease in the ß-galactosidase activity of the fusion. This last result was confirmed using the P_tet-phoP-lacZ fusion (repression of 3.5-fold,Figure 1B), indicating that, as shown previously for MicA[24], GcvB most likely acts at the post-transcriptional level. Importantly, this repressor effect of GcvB was also visible when GcvB was expressed from the chromosome; a deletion of thegcvB gene was sufficient to increase expression ofphoP-lacZ by 1.9-fold (Figure 1C).

GcvB and MicA Act Independently onphoP

One possible explanation for these results is that GcvB regulates thephoP-lacZ fusion by controlling the synthesis and/or activity of a post-transcriptional regulator ofphoP. Since MicA is so far the only post-transcriptional regulator ofphoP known to affect ourphoP-lacZ fusion, we analyzed the effect of GcvB onphoP expression in the absence of MicA. In this context, overproduction of MicA and GcvB from a plasmid caused a 3.4- and 4.3-fold decrease respectively in the activity of thephoP-lacZ fusion (Figure 2A), which is similar to what was observed inmicA⁺ cells. Consistent with this observation, deletion ofgcvB resulted in a 1.7- or 2.3-fold activation ofphoP-lacZ in wt ormicA⁻ cells respectively (Figure 2B). Therefore, GcvB acts independently of MicA to regulatephoP expression. In this experiment, deletion ofmicA has no significant effect on the expression ofphoP, because transcription of MicA is dependent on the RpoE sigma factor, which is not activated under the experimental conditions ofFigure 2B.

We had previously constructed a mutant form ofphoP-lacZ (phoPmut-lacZ, where the 4 nts directly downstream of the AUG start codon are changed from CGCG to GCGC,Figure 3A) such that this fusion is no longer controlled by MicA[24]. Interestingly, this mutant fusion is still controlled by GcvB, since its expression is up-regulated by 2.2-fold in aΔgcvB strain (Figure 2B, two last bars). This result suggests that the precise regions of thephoP mRNA that are required for MicA or GcvB action are different. Thus, GcvB acts onphoP, independently and apparently at a different site from that of MicA.

The R1 Region of GcvB Is Not Required for the Control ofphoP

GcvB is a pleiotropic regulator, whose expression is highest in exponentially growing cells in rich medium as a result of its control by the GcvA transcriptional regulator. GcvB directly regulates more than 20 genes, the large majority of which are targeted via a very well conserved single stranded G/U rich region of GcvB, referred to as R1 (Figure 3A). Even though this might be partially due to an experimental bias given that the R1 region was used as a “bait” in a bioinformatic search for targets, this region is clearly required for the control of almost all targets identified so far[13]. Since sRNAs often regulate multiple targets via a single conserved region[18],[29],[30], we reasoned that the R1 region was also likely to be involved in the control ofphoP, regardless of whetherphoP was a direct or indirect target. We therefore measured the expression of the P_BAD-phoP-lacZ fusion in the presence of a plasmid overexpressing either GcvB wt or a GcvB mutant in the middle of the R1 region (GcvBmutR1, seeFigure 3B). In this experiment, the chromosomal copy ofgcvB is deleted and the steady-state level of GcvBmutR1 was slightly lower than that of GcvB wt. Somewhat surprisingly, these two forms of GcvB repressed the expression ofphoP-lacZ to a similar extent (Figure 3C, left panel); in contrast, a previously identified target of GcvB R1 region,livJ, was, as expected, less regulated by GcvBmutR1 than by GcvB wt (Figure 3D). While this does not completely rule out a possible role for the R1 region in the control ofphoP (for instance, if pairing involves nts of R1 that are not affected by the mutR1 change or if alternative pairing(s) can take place with this mutant), this suggests that the role of R1 inphoP control is not as crucial as for the other targets of GcvB.

GcvB Is a Direct Regulator ofphoPQ

Because GcvB action onphoP was independent of MicA (see above), we next envisioned the possibility that it could directly pair withphoP to control its expression. If this interaction exists, we expected it to involve a region of GcvB outside of the R1. The TargetRNA program[31] was used to predict a potential pairing between GcvB and thephoP mRNA fragment encompassing nts −36 to +30 relative to the AUG (i.e. the region ofphoP that is present on thephoP-lacZ fusion). According to this prediction (Figure 3B), the region between nts 148 and 174 of GcvB can imperfectly pair withphoPQ mRNA in the translation initiation region (TIR), which is the most frequent binding site for negatively acting sRNAs. Interestingly, this corresponds to a region of GcvB that was shown to be mostly single-stranded in solution[12] and is now referred to as region R3. The relevance of this putative direct interaction was testedin vivo by compensatory changes. While the P_BAD-phoP-lacZ fusion was repressed by more than 4-fold upon overproduction of GcvB wt, this was not the case with the GcvBmutR3 variant, where nts 154 to 158 were changed from CUGUC to GACAG. Rather, expression of the fusion was increased by more than 2-fold (Figure 3C). The inability of GcvBmutR3 to repressphoP-lacZ expression is not due to an intrinsic instability, since it accumulates to a level similar to that of wt GcvB (Figure 3C). A possible explanation for the fact that GcvBmutR3 activatesphoP-lacZ is that it could titrate Hfq when overexpressed, leading to changes in expression of Hfq-regulated genes, such asphoP (see[28],[32] for examples of competition for Hfq). In contrast, GcvBmutR3, but not wt GcvB, caused an 8-fold decrease in the activity of the compensatory mutant fusion (Figure 3C, right panel), clearly showing that GcvB andphoP mRNA directly interactin vivo. It is also worth noting that MicA efficiently repressed both the wt and the mutant fusion, which confirms that GcvB and MicA pair at different loci ofphoPQ mRNA. In addition, when mutations in the R1 and R3 regions of GcvB were combined, the resulting GcvBmutR1R3 repressed the expression ofphoPmutR3-lacZ, but not that ofphoP-lacZ (Figure S1A). This again indicates that, at least when GcvB is overexpressed, its R1 region is not involved in the control ofphoP.

phoP mRNA InteractsIn Vitro with MicA and GcvB

To provide experimental support to the proposedphoP-MicA andphoP-GcvB base-pairing interactions (Figure 3B), a structural probing analysis ofphoP mRNA alone or in the presence of either sRNA was performedin vitro using chemical probes (Figure 4, A and B). DMS (dimethyl sulfate), CMCT (1-cyclohexyl-3-(2-morpholinoethyl)carbodiimide metho-p-toluene sulfonate) and kethoxal (1-1-dihydroxy-3-ethoxy-2-butanone) respectively modify unpaired adenosine (and to a much lesser extent cytidine), uridine and guanosine residues. According to our probing data, the secondary structure of the 5′ region ofphoP mRNA appears as a long irregular stem-loop which is hold by seven double-stranded elements named H1 to H7, separated by bulges or loops (Figure 4C). Upon addition of MicA, most nts from positions −13 to +11 ofphoP mRNA, which include the nts forming the 5′ strands of H4, H5 and H6, display either a decreased reactivity towards the probes or correspond to RT-stops or -pauses which occur in a region rich in GC pairs (Figure 4, A and D). This model is also consistent with the fact that many nts located between positions +27 to +45 ofphoP mRNA, which include all the nts forming the 3′ strands of H4, H5 and H6 in the absence of MicA, become more reactive in the presence of MicA (Figure 4, A and D). In conclusion, the interaction betweenphoP mRNA and MicA relies on (i) the disruption of at least three of these elements, namely H4, H5 and H6, and (ii) the formation of an extended base-pairing interaction between nts −15 to +11 ofphoP mRNA and nts 4 to 31 of MicA, whereby both the Shine-Dalgarno (SD) sequence and thephoP translation start codon are base-paired (Figure 4D). Upon addition of MicA, further reactivity enhancements inphoP mRNA nts are observed outside of the proposedphoP mRNA-MicA duplex (see nts +53 and +54 in H1, −21 and +46 to +49 in H3, −16 which joins H3 to the duplex, +14 to +21 in H7 and its apical loop,Figure 4D). It is likely that these changes are due to either local breathing or even disruption of H1, H3 and H7, which are destabilized by the binding of MicA. Also, decreased reactivities are observed (see nts +12 which joins the duplex to H7 and +23 in H7,Figure 4D) for which we have no explanation.

In contrast, the duplex formed byphoP mRNA and GcvB seems shorter as it requires only the disruption of H3 and H4 to form; it is centered around and blocks the SD sequence, which is in complete agreement with thein vivo data. Indeed, nts displaying decreased reactivities towards the probes or corresponding to a region where GcvB-induced RT-stops or -pauses occur are clustered between nts −17 and −11 ofphoP mRNA (Figure 4B and 4E). However, the duplex is likely to be subject to breathing as a certain number of nts located on both side of the cluster become more reactive in the presence of GcvB (see nts −21 to −19, −10 and −9,Figure 4E). Additional reactivity enhancements have been mapped outside of the proposed duplex (see nts −5, +2, +3, +5 and +31 to +36 in H5 and H6 and in the bubble located in between,Figure 4E), which are probably due to breathing of H5 which results from its destabilization by the binding of GcvB. Other regions ofphoP mRNA located outside of the predicted duplex are subject to increase or decrease in reactivity in the presence of GcvB (see nts located in H6 and H7 and in the bubble in between,Figure 4E). They can be due to some rearrangement of the overall structure ofphoP mRNA upon GcvB-binding and/or to a supplementary interaction betweenphoP mRNA and GcvB. For instance, nts +9 to +17 ofphoP mRNA, several of which appear protected upon GcvB-binding, could theoretically pair with nts 89 to 97 of GcvB. While ourin vivo data show that this putative supplementary interaction is not sufficient for control, it could nevertheless play a role in stabilizing thephoP mRNA-GcvB duplex or in increasing the kinetics of association. At this stage, its existence and importance remains to be experimentally addressed. Finally, the reactivities of nts +37 to +50, which form the 3′ strands of H3 and H4 in the absence of GcvB, could not be assessed with confidence because of the presence of several RT-stops or –pauses which are also present whenphoP mRNA alone is reverse-transcribed in the absence of the probe (data not shown).

Both MicA and GcvB Decrease the Level ofphoPQ mRNA

Target-mRNAs of negatively acting Hfq-dependent sRNAs are frequently degraded upon sRNA production. Therefore, to confirm the results obtained above by gene fusion, the levels ofphoPQ mRNA were analyzed by Northern-Blot upon overexpression of MicA, GcvB or their mutant derivatives, using a chromosomal P_BAD-phoPQ construct (Figure 5A). In this experiment, transcription of thephoPQ operon is again expected to start 36 nts upstream of thephoP start codon and has been put under the control of the P_BAD promoter for two reasons: (i) to focus only on promoter-independent regulation and (ii) because of the low abundance of thephoPQ mRNA when expressed from its own promoters under the experimental conditions used here. With this construct, several specific bands are visible. The upper band migrates below a 3 kb RNA marker and most likely corresponds to the wholephoPQ mRNA, while the bands of lower molecular weight could result from either alternative transcription or processing events (Figure 5A). MicA, GcvB and GcvBmutR1 induce a decrease inphoPQ mRNA levels, but not MicAmut and GcvBmutR3, that have lost the ability to controlphoP expression. This is in complete agreement with the results obtained with thephoP-lacZ fusions. Pairing of MicA and GcvB to thephoPQ mRNA is therefore likely to induce a degradation of this mRNA.

MicA and GcvB Differentially Affect the Level of PhoP Protein

The effect of MicA or GcvB on the steady-state levels of the PhoP protein was then investigated by Western-Blot analysis, in a strain wherephoPQ is expressed from its own promoter. As expected, MicA overexpression resulted in a strong decrease in the amount of PhoP (Figure 5B), while overexpression of MicAmut had no noticeable effect. When GcvB was overexpressed, PhoP levels were decreased, albeit to a much lesser extent than upon MicA overproduction. This is rather surprising since pMicA and pGcvB had a similar effect on the expression ofphoP-lacZ (Figure 1,Figure 2, andFigure 3) and on the levels ofphoPQ mRNA (Figure 5A). Interestingly, pGcvBmutR1, whose effect was also similar to that of pMicA and pGcvB in the previous experiments, is more efficient than pGcvB in down-regulating the levels of PhoP protein. Finally, GcvBmutR3 overproduction had no effect on the levels of PhoP, which is consistent with its inability to repressphoP expression (Figure 5B).

Therefore, control ofphoP by GcvBmutR1 or MicA results, as expected, in a clear decrease of the PhoP protein levels. Surprisingly however, this decrease is only modest with wt GcvB, most likely because its R1 region has pleiotropic effects in the cell under the conditions used here, as discussed below.

MicA and GcvB Differentially Affect Expression of the PhoP Regulon

We then tested whether MicA and GcvB can control the expression of the PhoP regulon by repressing PhoQ-PhoP synthesis. For this purpose, the expression of 4 genes whose transcription is directly activated by PhoP[25] was analyzed under conditions where either MicA or GcvB was overproduced (Figure 6). These 4 genes areompT,mgtA,yneM andmgrR, that encode an outer membrane protease, a magnesium transporter, a protein of unknown function located in the outer membrane and a sRNA regulator of LPS modification respectively. As expected, MicA induced a >2.5-fold decrease in expression of these 4 targets as analyzed by either translational fusions (forompT,mgtA) or by transcriptional fusions (formgrR andyneM) to (Figure 6A). This decrease is most likely due tophoP regulation, since MicAmut, that does not regulatephoP, does not affect expression of these target genes. Similarly, the overproduction of GcvBmutR1, that also repressesphoP, led to a ∼2-fold decrease in the expression of the 4 fusions, while overexpression of GcvBmutR3 does not, in agreement with its inability to controlphoP. Finally, when the same experiment was carried out in the GcvB overproducing strain, no decrease was observed in the expression of the 4 members of the PhoP regulon that were tested. Instead, activity ofmgrR- andompT-lacZ was unchanged, while activity ofmgtA- andyneM-lacZ was increased by 1.6 and 2.1-fold respectively (Figure 6A). These results obtained by gene fusion were confirmed when the levels ofompT or MgrR RNAs were analyzed by Northern-Blot (Figure 6B). Indeed, MicA and GcvBmutR1 induced a decrease in the level of both RNAs, whereas MicAmut, GcvB wt and GcvBmutR3 did not.

Therefore, MicA and GcvBmutR1 repress the PhoP regulon, by controlling expression ofphoP. However, wt GcvB does not, which is consistent with the only modest decrease observed in PhoP levels upon its overproduction.

MicA and GcvB Inhibit Ribosome Binding to thephoP TIR

In most cases, negatively acting sRNAs base-pair with their target-mRNAs in the TIR and occlude the RBS, thereby preventing ribosome binding and translation initiation. This is frequently accompanied by a degradation of the target-mRNA, possibly as a consequence of translational block, or in a process that is directly induced by the sRNA pairing to the target-mRNA. Since MicA and GcvB both pair tophoPQ mRNA in the TIR of the first cistron and decreasephoPQ mRNA levels, toeprinting experiments were performed in order to determine whether they also inhibited ribosome binding. In these experiments, addition of 30S ribosomal subunit and initiator tRNA to a ∼200-ntphoP mRNA fragment transcribedin vitro induced an arrest of reverse-transcription, that was visible on a sequencing gel as a band at position +16, a classical toeprint position (Figure 7, A and B, lanes 3 and 2 respectively). When increasing concentrations of MicA were incubated withphoP mRNA prior to the addition of 30S and fMet-tRNA, the intensity of this band progressively decreased (Figure 7A, lanes 4–6). This was not observed when equal amounts of MicAmut were added instead (lanes 8–10), suggesting that it is the pairing of MicA tophoP TIR that inhibits ribosome binding. Similar results were observed with GcvB, whose addition inhibited the appearance of the +16 toeprint, even more efficiently than MicA (Figure 7B, lanes 7–8). Again, this is most likely due to the pairing between GcvB andphoP since GcvBmutR1, that should still pair withphoP, also inhibited the toeprint, while GcvBmutR3 and GcvBmutR1R3, that should not bind tophoP, inhibited the toeprint much less efficiently (Figure 7B, lanes 11–12, andFigure S1B). It is interesting that GcvB is much more efficient than MicA in inhibiting toeprint, and that GcvBmutR3 still inhibits toeprint to some extent. This might be related to the existence of a bipartite interaction betweenphoP and GcvB (see above).

Furthermore, as already observed in probing experiments (Figure 4), addition of MicA, but not MicAmut, tophoP mRNA in the absence of 30S subunits also induced stops or pauses of reverse-transcription, as indicated by the bands at positions +6 to +8 (Figure 7A, lanes 2, 4, 5 and 6). This corresponds to the 3′ end of the duplex between MicA andphoP (Figures 3A,4D,[24]), indicating that this duplex is stable enough to induce pauses in reverse-transcription. In contrast, while GcvB pairs withphoP in vitro, given the results of the probing experiments and the toeprint inhibition, its binding does not induce pauses or stops of the reverse transcription that are sufficiently strong to be observed in this experiment. This is in contrast to what was observed inFigure 4 and this discrepancy is most likely due to the use of different experimental conditions in the probing and toeprint experiments. In fact, under conditions where the signal is highly amplified, reverse transcriptase stops are visible in the toeprint experiments (data not shown). Hfq protein was not included in thesein vitro assays, because of the risk of non-specific interactions with RNA. The fact that, even in the absence of this chaperone, MicA and GcvB could both pair tophoP mRNAin vitro (i.e. in the absence of RNases) suggests that the requirement for Hfqin vivo is, at least in part, explained by its ability to protect MicA and GcvB from degradation.

In summary, both MicA and GcvB inhibit ribosome binding by pairing tophoPQ TIR. This translational block could be the step leading to the degradation of the target-mRNA in presence of the regulatory sRNAs observedin vivo.

GcvB as a Novel Regulator ofphoP Expression: A Control Likely Conserved in Several Enterobacteria

In this study, we identifyphoPQ mRNA as a new target of theE. coli GcvB sRNA. After MicA, this is thus the second sRNA regulator of this operon. Similar to MicA, GcvB directly controlsphoPQ expression by pairing to the TIR ofphoP, although at sequences slightly different from those of MicA. These pairings cause a steric inhibition of ribosome binding as seen by toeprint experiments and, possibly as a consequence of this translational control, induce degradation of thephoPQ mRNA. Furthermore, this work identifies a novel pairing region of GcvB, namely R3, essential forphoP control. Interestingly, this region was predicted in a computational approach as a potential target-binding region of GcvB (together with the R1) on the basis of its conservation and accessibility[33], and was proposed to participate in the control ofcycA expression[34]. Whether GcvB controls yet additional genes through its R3 region remains to be investigated.

This region R3 is with R1 and R2 one of the most conserved in GcvB among enterobacteria (Figure S2A). However,phoP was not identified as a GcvB target inSalmonella in a recent study combining microarray analysis following GcvB pulse-expression and bioinformatic prediction based on complementarity to the R1 region[13]. While this could be due to the low abundance ofphoPQ mRNA and to the fact that this control does not rely on R1, this could also indicate that the control ofphoP by GcvB that exists inE. coli is not conserved inS. typhimurium. Consistent with this, predictions of pairing between GcvB R3 region and the TIR ofphoP mRNA inSalmonella identified only 4 consecutive complementary nts at the most, which is probably too short to ensure specific binding. Furthermore, a preliminary analysis of potential interactions between GcvB R3 and thephoP TIR in different families of enterobacteriaceae suggests that GcvB could controlphoP in species such asKlebsiella pneumoniae,Photorhabdus luminescens,Proteus mirabilis,Serratia proteamaculans,Shigella flexneri,Xenorhabdus bovienii andRahnella (Figure S2B).

Why Does wt GcvB, but Not GcvBmutR1, Only Modestly Affect PhoP Protein Levels?

At first glance, it is quite surprising that, even though MicA, GcvB and GcvBmutR1 similarly repressphoP expression when followed by gene fusion tolacZ or mRNA levels, their effects on the PhoP protein are quite different. Indeed, while MicA and GcvBmutR1 strongly decreased the cellular level of PhoP, as expected, the effect of wt GcvB was much more moderate (Figure 5B). One noticeable difference in those experiments is that when the PhoP protein levels were assessed,phoPQ was expressed from its own promoter. In contrast, in both the experiments with gene fusion or mRNA levels, its transcription is driven by an heterologous promoter (P_BAD), withphoPQ andphoP-lacZ mRNAs expected to originate at the same transcription start site than from P₁. Therefore, one could hypothesize that wt GcvB would activate PhoP synthesis from transcripts originating from promoters P₂ or P₃ (upstream of P₁), or by acting onphoPQ transcription, in addition to repress its expression post-transcriptionally. These possibilities were experimentally ruled out because (i) wt GcvB similarly repressed expression of aphoP-lacZ fusion whose 5′end is identical to the 5′end initiating from P₁ or from P₂ promoter (Figure S3A) and (ii) wt GcvB only poorly affectsompT expression (and even increasesyneM expression) in a strain where all thephoPQ promoters have been replaced by P_BAD (Figure S3B).

One could also envision that, when expressed from its own promoter under non-inducing conditions (as inFigure 5B),phoP expression is poorly affected by wt GcvB, whose R1 region can pair with other competing targets. This competition could not take place with GcvBmutR1, where the R1 region is mutated, in agreement with its ability to controlphoP in all experiments. In contrast, whenphoP expression increases, for instance because its transcription is driven by an induced P_BAD promoter, it would now become available for repression by wt GcvB, because it would outcompete other GcvB targets, hence the stronger effect of wt GcvB onphoP inFigures 3C and5A for instance. It will now be interesting to study how the induction ofphoP from its own promoter and the expression of GcvB R1 targets will impact the control ofphoP by GcvB; in other words, whether this model is physiologically relevant.

Yet another possibility to explain the difference betweenphoP-lacZ expression,phoPQ mRNA and the PhoP protein levels in presence of pGcvB is that, in addition to its negative effect onphoP expression at the translation initiation step, wt GcvB could stabilize the PhoP protein. Such a regulatory event would affect only PhoP levels, but not the activity ofphoP-lacZ orphoP mRNA levels. This putative stabilization would be dependent on the R1 region of GcvB and could be mediated by one or several of its targets. Furthermore, one could wonder whether this stabilization is related to the phosphorylation status of PhoP protein. Because these R1 targets that could directly or indirectly control PhoP are likely to be multiple, it might be difficult to identify them by a genetic approach. Again, it is tempting to speculate that the expression of these targets, as well as the availability of the R1 region to regulate them (dependent on the expression of all GcvB targets and their relative affinity for the sRNA), will play an important role in the control of the PhoP regulon by GcvB. While under our experimental conditions, the amount of PhoP protein is not strongly affected by wt GcvB, there might be conditions where it could be. This would provide a mechanism to establish a hierarchy among the different GcvB-targets and to monitor the regulatory outcomes of GcvB-mediated controls.

A Novel Example of the Complex Interplay between sRNAs and Transcriptional Regulation Networks

In a previous work, we showed that MicA, which is induced by envelope stress, repressed the expression ofphoPQ. This finding related the activity of this operon to the cell envelope status[24]. Our present findings show that GcvB relates amino acid (or peptide) uptake and metabolism to the expression ofphoPQ. The induction of GcvB occurs in the presence of two different amino acids with two different mechanisms. First through the GcvA/GcvR repressing complex that is inactivated in the presence of glycine, and second through the global regulator Lrp, whose repression is alleviated in the presence of leucine[9],[11] (Figure 8). Although theraison d'être of such a connection between amino acid uptake/metabolism and PhoQ/PhoP regulon activity is not obvious, such a relationship has already been observed with several targets of this regulon. For instance, expression ofmgtA andmgtCBR genes is derepressed under conditions of proline limitation due to the presence of a proline-rich open reading frame in their leader mRNAs[35],[36]. Another example is the proline transporter encoded by theproP gene that is also regulated by PhoQ/PhoP[37].

The complexity of the relationship between GcvB and thephoPQ regulon is highlighted by two experimental data. The first is the surprising way wt GcvB fails to strongly decrease PhoP levels as discussed above. The second is related to the recent results of a deep-sequencing study indicating that PhoP positively regulates GcvB levels in the cell[38](Figure 8). However, GcvB levels were unmodified when the magnesium concentration in the growth medium was varied. Therefore, GcvB could also modulate the degree of PhoQ/PhoP activation depending on the inducing signals.

Interestingly, there are already many examples of connections between sRNAs and TCS, as several sRNAs were previously shown to control TCS and conversely[39]. In addition, the negative feedback loop that exists betweenphoP and GcvB is reminiscent of other feedback loops involving sRNAs[40]. As in most cases however, the properties and possible advantages of this feedback loop in bacterial physiology remain to be experimentally addressed.

Our search for Hfq-dependent sRNAs regulators of the PhoQ/PhoP TCS was initially motivated by the fact thatphoP expression was up-regulated in anhfq mutant strain independently of MicA. Interestingly however, even though GcvB is partially responsible for this Hfq-effect, expression ofphoP is still higher in anhfq mutant than inhfq⁺ cells in the absence of both MicA and GcvB (data not shown). This suggests that there might be even more sRNAs controlling PhoQ/PhoP, which would allow the integration of yet additional signals to fine-tune expression of this central TCS.

General Microbiological Techniques and Strain Construction

Strains and plasmids used in this study are listed inTable 1, and sequences of the oligonucleotides inTable S1. Strains were grown aerobically in LB medium at 37°C. When needed, antibiotics were used at the following concentrations: ampicillin 150 µg/ml, tetracyclin 10 µg/ml, kanamycin 25 µg/ml or chloramphenicol 10 µg/ml. PCR amplification was performed using the Phusion DNA polymerase (New England Biolabs). IPTG (isopropyl-ß-D-thiogalactopyranoside) was used at a final concentration of 100 µM.

Replacement ofgcvB gene by a kanamycin or tetracyclin resistance cassette was engineered by recombineering of a cassette amplified by PCR (with ΔgcvB::kanfor and rev, or ΔgcvB::tetfor and rev oligonucleotides) and flanked by homology regions upstream and downstream ofgcvB into a strain carrying a mini-lambda allowing recombineering upon induction, such as NM300 or NM1200 for instance. These mutant alleles, as well asΔmicA::tet[24] orphoP::kan[41] were then moved by P1 transduction when necessary.

Strains carrying gene fusions tolacZ were either obtained from different sources or constructed in this study by recombineering into strain PM1205[19] or strain MG1508. In strain MG1508, acat-sacB cassette following the P_LtetO-1 promoter[42] is placed upstream of thelacZ gene in an MG1655 derivative that carries a mini-lambda.

Strain MG1173 carries a translationalompT-lacZ fusion, whose construction was described previously, at the lambda attachment site[18]. Strains KM112 and KM194 were also described elsewhere[43]. They contain respectively the promoter region up to nt+10 of MgrR, or the promoter region ofyneM up to nt+91 (nt+1 is the transcriptional start site) upstream oflacZ chromosomal gene, starting 17 nts upstream of its ATG start codon. The translational P_tet-phoP-lacZ fusion was constructed by replacing thecat-sacB cassette of the P_tet-cat-sacB-lacZ construct in strain MG1508 with a PCR fragment encompassing nts −36 to +30 (relative to the ATG start codon) ofphoP between homology regions to P_tet andlacZ respectively. This PCR fragment was generated with primers 5′Ptet-phoP and 3′phoP-lacZ. Recombinants were selected on LB-agar plates without NaCl supplemented with 6% sucrose, and further verified as in[24]. Similarly, construction of the P₁-mgtA-lacZ fusion was done by recombineering of a PCR fragment carrying nts −330 to +30 ofmgtA in MG1508, except that the homology regions were upstream of Ptet and withinlacZ (see primers 5′P₁mgtA and 3′mgtA-lacZ).

For strains MG1510 (P_BAD-phoPmutR3-lacZ) and AC0067 (P_BAD-livJ-lacZ), PCR fragments were generated with primers phoPmutR3for and 3′phoP-lacZ, or 5′LivJ-lac and 3′LivJ-lac respectively, then recombined into strain PM1205.

Construction of strains where thephoPQ operon, wt or interrupted by a kanamycin resistance cassette, is expressed from a P_BAD promoter was as follows. First, the chloramphenicol resistance cassette followed by the P_BAD promoter was amplified by PCR from plasmid pTM26[44] with primers 5′Cm-PBAD-phoPQ and 3′Cm-PBAD-phoPQ. This product was then recombined in strain NM300 (or a derivative carrying thephoP::kan allele moved by P1 transduction using AB043[41] as the donor strain). After selection on LB-chloramphenicol plates, the P_BAD promoter and beginning ofphoP gene were checked by sequencing, and resistance to kanamycin was verified for the P_BAD-phoP::kan construct. These alleles were then moved by P1 transduction using DJ624 as the recipient strain and selecting for chloramphenicol resistant clones to create strains MG1516 and MG1517. MG1517 was checked for resistance to kanamycin.

Plasmids

ß-Galactosidase Assay

Overnight cultures were diluted 500 fold in fresh medium (see below for exact medium composition) and grown to mid-exponential phase (OD at 600 nm∼0.4). The ß-galactosidase activity was then measured as in[45] and expressed in Miller units. Alternatively, the activity was measured in a 96-wells plate for some experiments. In this case, 100 µl of cells were mixed with 50 µl of permeabilization buffer containing 200 µg/ml of polymixin B[46]. After addition of 50 µl of ONPG, the absorbance at 420 nm was followed over-time and the ß-galactosidase activity was calculated as the slope of the resulting curve. It is expressed in arbitrary units.

Cells were grown in the following media: forFigure 1A, LB-Ampicillin-IPTG-Arabinose 0.002% or 0.02%; forFigure 1B, LB-Ampicillin-IPTG; forFigure 1C, LB; forFigure 2A and3C, LB-Ampicillin-IPTG-Arabinose 0.02%; forFigure 2B, LB-Arabinose 0.002%; forFigure 3B, LB-Ampicillin-IPTG-Arabinose 0.02%, but IPTG was not included in the overnight cultures; forFigure 6A, LB-Tetracyclin-IPTG; forFigure S1, LB-Tetracyclin-IPTG-Arabinose 0.02%.

Values of ß-galactosidase activity given in the paper are the average of at least two independent experiments and are listed inTable S2.

RNA Extraction and Northern Blot Analysis

RNA was extracted following the hot phenol method as previously described[47] and using 650 µl of cells. For experiment ofFigure 3, RNA was extracted at the same time than samples were taken to measure ß-galactosidase activity. For experiments ofFigures 5 and6, cells were grown overnight in LB-Tetracyclin-IPTG-Arabinose (0.2% forFigure 5 and 0.02% forFigure 6), then diluted in fresh medium and grown to mid-exponential phase (A₆₀₀∼0.4 forFigure 6B) or to stationary phase (A₆₀₀∼2 forFigure 5) before RNA was extracted. For Northern analysis, a constant amount of RNA was separated on an 8% acrylamide TBE-urea gel (for MicA, GcvB, MgrR and SsrA RNAs) or on a 1% denaturing agarose gel (forompT andphoP mRNAs) and transferred to an Hybond-N+ membrane. Detection was then performed using biotinylated probes and the Ambion brightstar detection kit following manufacturer's instructions.

Western Blot Analysis

Overnight cultures in LB-Tetracyclin-IPTG of strain MG1173 (wt) or MG1446 (phoP⁻), transformed with pBRplac and derivatives, were diluted 500-fold in the same medium and grown to mid-exponential phase. Cells were then pelleted and resuspended in SDS-sample buffer with DTT (New England Biolabs) at a final concentration of 15 OD₆₀₀/ml. These samples were then boiled for 5 minutes and 15 µl were loaded on a 15% SDS-PAGE gel. Proteins were then transferred to an Hybond-C super membrane (Amersham) and the PhoP protein was detected using a 1∶1000 dilution of an anti-PhoP antiserum (from Mark Goulian) and the Immun-Star WesternC Chemiluminescent Kit (Biorad). EF-Tu was immunodetected from the same membrane. A representative blot from three independent experiments is shown.

In Vitro RNA Transcription

PCR templates for thein vitro transcription of MicA or GcvB (and their derivatives) were prepared from the pBRplacMicA(mut) or pBRplacGcvB(mutR1, mutR3 or mutR1R3) plasmids, using the oligonucleotides 5′T7MicA(mut) and 3′T7MicA, or 5′T7GcvB and 3′T7GcvB respectively. Note that one or two G residues are added at the 5′end of MicA and GcvB respectively. ForphoP, a PCR fragment corresponding to nts −36 to +169 ofphoP mRNA preceded by two G residues was amplified from genomic DNA using oligonucleotides 5′T7phoP and 3′T7phoP. After purification, these PCR products were used inin vitro transcriptions reactions with the T7 RNA polymerase of Stratagene (forphoP) or the T7 Megascript kit from Ambion (for MicA and GcvB) following manufacturer's instructions. After phenol extraction and precipitation with ammonium acetate, RNA were purified using G-50 Microspin columns (GE Healthcare).

Toeprinting Assays

Toeprinting assays were adapted from Hartzet al.[48] as follows. 0.5 pmol. ofphoP transcript were incubated with 2 pmol. of phoP-Cy5-probe#1, an oligonucleotide complementary to nts 52 to 71 ofphoP ORF and labeled with a Cy5 group at its 5′ end, in a buffer containing 10 mM Tris-acetate pH 7.4, 60 mM ammonium chloride and 6 mM ß-mercaptoethanol. When required, sRNAs were added to this mix at the desired concentrations. These mixtures were denatured by heating at 80°C for 3 minutes, followed by a rapid cooling in an ethanol/solid CO₂ mix. They were then thawed on ice and magnesium was added at a final concentration of 10 mM. For experiment ofFigure 7A, an additional incubation step at 37°C for 10 minutes was performed at this stage. 190 µM dNTPs and 2.5 µM of initiator tRNA^fMet were added together with 0.5 µM 30S subunits, and the mixtures were incubated at 37°C for 10 minutes. 1 unit of AMV RT (Finnzymes) was then added and cDNA synthesis was performed at 37°C for 20 minutes. Reactions were stopped by addition of formamide and EDTA, analyzed on a 6% sequencing gel together with sequencing reactions and RT stops were visualized using a typhoon fluorescent scanner set up for Cy5 detection.

Structure Probing and Protections

1.25 pmol. ofphoP transcript, mixed in water with 6.25 pmol. of sRNA when required, were denatured as above. After a slow thaw-out on ice, samples were incubated at 37°C for 10 minutes in order to allow sRNA andphoP message to pair. Samples were then diluted in a buffer containing 50 mM sodium cacodylate pH 7.5, 10 mM magnesium acetate and 50 mM ammonium chloride prior to DMS treatment, or in the same buffer except that sodium cacodylate is replaced by sodium borate pH 8.0 prior to CMCT or Kethoxal treatment. After a 10 minutes incubation at 25°C, 1 µg ofL. lactis 23S rRNA was added, followed by 0.1 volume of DMS or kethoxal (stock solutions are 1/30 in ethanol or at 4 mg/ml in 20% ethanol respectively) and samples were incubated at 25°C for 5 minutes. For CMCT treatment, 0.1 volume of 100 mg/ml CMCT in the previous buffer containing sodium cacodylate was added and samples were incubated at 25°C for 10 minutes. Modified RNA were then precipitated with ammonium acetate (and 100 mM sodium borate pH 8.0 for samples treated with Kethoxal) and resuspended in water (or in 12.5 mM sodium borate pH 8.0 for samples treated with Kethoxal).

For reverse transcription, phoP-Cy5-probe#2, complementary to nts 87 to 106 of phoP ORF, was added to those samples at the final concentration of 1 µM. This was followed by the addition of 2 units of AMV RT (Finnzymes), together with 1 mM dNTPs and 4 mM DTT. cDNA synthesis and analysis was then performed as described above.

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Supplementary Materials