Tn5 transposition in Escherichia coli is repressed by Hfq and activated by over-expression of the small non-coding RNA SgrS
© Ross et al.; licensee BioMed Central Ltd. 2014
Received: 18 August 2014
Accepted: 11 November 2014
Published: 30 November 2014
Hfq functions in post-transcriptional gene regulation in a wide range of bacteria, usually by promoting base pairing of mRNAs with trans-encoded sRNAs. It was previously shown that Hfq down-regulates Tn10 transposition by inhibiting IS10 transposase expression at the post-transcriptional level. This provided the first example of Hfq playing a role in DNA transposition and led us to ask if a related transposon, Tn5, is similarly regulated.
We show that Hfq strongly suppresses Tn5 transposition in Escherichia coli by inhibiting IS50 transposase expression. However, in contrast to the situation for Tn10, Hfq primarily inhibits IS50 transposase transcription. As Hfq does not typically function directly in transcription, we searched for a transcription factor that also down-regulated IS50 transposase transcription and is itself under Hfq control. We show that Crp (cyclic AMP receptor protein) fits these criteria as: (1) disruption of the crp gene led to an increase in IS50 transposase expression and the magnitude of this increase was comparable to that observed for an hfq disruption; and (2) Crp expression decreased in hfq − . We also demonstrate that IS50 transposase expression and Tn5 transposition are induced by over-expression of the sRNA SgrS and link this response to glucose limitation.
Tn5 transposition is negatively regulated by Hfq primarily through inhibition of IS50 transposase transcription. Preliminary results support the possibility that this regulation is mediated through Crp. We also provide evidence that glucose limitation activates IS50 transposase transcription and transposition.
KeywordsTn5/IS50 Hfq Crp SgrS DNA transposition
Transposase proteins catalyze the chemical steps in bacterial transposition reactions. It follows that the regulation of expression of these genes is a critical feature in dictating the transposition frequency of most transposons. In many instances, including Tn10/IS10 and Tn5/IS50, transposase gene promoters are inherently weak. In addition, DNA adenine methylase (DAM) limits initiation of IS10 and IS50 transposase gene transcription by methylating promoter elements ,. These factors together make transcription initiation a limiting step in Tn10/IS10 and Tn5/IS50 transposition reactions ,. There are also examples where translation of transposase transcripts is subject to both intrinsic and host levels of regulation. In the case of IS10 transposase, the ribosome binding site is inherently weak and the transposon encodes an antisense RNA that binds the translation initiation region (TIR), blocking ribosome binding ,. There is also evidence that the ‘host’ protein Hfq helps mediate the pairing interaction between the antisense RNA and the IS10 transposase transcript ,.
As noted above, Hfq has been implicated in the regulation of Tn10/IS10 transposition. Under conditions of hfq deficiency, a large increase in both Tn10/IS10 transposition (up to 80-fold) and transposase expression (up to 7-fold) were observed. The existing evidence is consistent with Hfq acting as a negative regulator of IS10 transposase expression by both antisense dependent and independent pathways. In support of the latter, it was found that hfq deficiency (or hfq − ) had a significant impact on Tn10 transposition even when the level of antisense RNA was insufficient to impact on transposase expression (that is when Tn10 is present in single copy in the bacterial chromosome). In addition, there was a synergistic increase in transposase expression when both hfq and the antisense RNA were knocked out, implying that Hfq does not function exclusively in the same pathway as the antisense RNA .
Taking the above results into account, and considering that most bacterial transposition systems are not regulated by antisense RNAs, we wondered if Hfq might play a more general role in regulating transposition systems. In the current work, we tested this hypothesis by asking if Tn5 transposition is also regulated by Hfq. Like Tn10, Tn5 is a composite transposon (Figure 1A). The two transposons are closely related but Tn5 lacks an antisense RNA regulatory system and consequently if Hfq were to regulate this system at the post-transcriptional level, it is likely that a trans-encoded sRNA would play a role -. Tn5 does encode an inhibitor protein that limits Tn5/IS50 transposition by dimerizing with the transposase protein, forming an inactive complex . Transposase and the inhibitor protein are expressed from overlapping promoters, P1 and P2 (color coded in Figure 1A), with the inhibitor transcript (T2) being expressed at a higher level than the transposase transcript (T1). T1 expression is down-regulated by DAM (reviewed in ). There is some evidence that P1 is also negatively regulated by LexA, an SOS-inducible transcriptional repressor . However, there is little else known with regard to host proteins that influence either transposase transcription or translation.
In the current work, we show that both Tn5 transposition and IS50 transposase expression increase significantly in E. coli under conditions of hfq deficiency. However, unlike the situation in Tn10/IS10 transposition, the up-regulation of IS50 transposase expression appears mainly to be due to an increase in transposase gene transcription. As Hfq does not typically function directly in transcription, we looked at the possibility that Hfq regulates IS50 transposase expression by controlling the expression of a transcription factor. Towards this end, we provide evidence that Hfq acts in a regulatory network with Crp (cyclic AMP receptor protein) to down-regulate IS50 transposase transcription. Finally, we demonstrate that over-expression of an sRNA (SgrS) activates expression of the IS50 transposase gene specifically when cells are grown with glucose as the sole carbon source. Evidence is presented that this up-regulation is a consequence of glucose limitation, demonstrating that the IS50 transposase promoter (and Tn5 transposition) is responsive to the nutrient status of the cell.
Hfq is a potent negative regulator of Tn5 transposition
We also performed a complementation assay in the DBH179 strain background to further test that the increase in transposition reported above in hfq− was actually due to the absence of Hfq, as opposed to possible polar effects of the hfq disruption allele. Towards this end, we introduced hfq on a low-copy plasmid (pDH700) into the hfq− strain and measured Tn5 transposition as above. We observed nearly complete complementation by plasmid-borne hfq, as transposition was reduced approximately 45-fold relative to when no hfq was present (Figure 2A). Furthermore, plasmid-encoded variants of hfq, including K56A and Y25A, which are impaired for RNA-binding at the ‘proximal’ and ‘distal’ surface, respectively, failed to complement hfq deficiency . This confirms that specific functions of Hfq, namely interaction with RNA via known RNA-binding surfaces, are required for effective repression of Tn5 transposition.
We also tested the impact of hfq deficiency on Tn5 transposition in a second donor strain background (DBH261) via the ‘mating out’ assay (Figure 2B). In this experiment hfq− also caused an increase in Tn5 transposition, although the magnitude of the effect was smaller (approximately 9-fold) than reported for the DBH179 strain background.
IS50 transposase expression increases in hfq − cells
Hfq impacts steady-state levels of full-length IS50 transposase mRNA
We also looked at the combined impact of knocking out Hfq and blocking DAM methylation on T1 levels (lanes 19 to 23 in Figure 5A). In comparison to wt, the ‘double mutant’ situation resulted in a 45-fold increase in T1 levels. Based on the observed synergy, we think it unlikely that the observed impact of deleting hfq is linked to the regulation of dam expression.
Taken together, the results from Figures 3, 4, 5 and 6 show that IS50 transposase expression is substantially reduced in an hfq + relative to an hfq − strain and that hfq status primarily affects transposase transcription.
Regulation of Tn5 transposase expression by global transcriptional regulators
We next asked if Crp expression was regulated by Hfq. Notably, work done in Yersinia pestis has shown that Hfq positively regulates Crp expression at the post-transcriptional level . Towards this end we performed Western blot analysis with a Crp antibody on E. coli cell extracts from wt (DBH303), hfq − (DBH306) and crp − (DBH307) strains (Figure 7C). The results show that lower levels of Crp are present in the hfq − strain, which is consistent with Hfq also being a positive regulator of Crp expression in E. coli.
Finally, we assessed the impact of knocking out crp on Tn5 transposition frequency using the ‘mating out’ assay (Figure 7D). In the absence of crp, Tn5 transposition increased 7-fold, which is consistent with results from the transposase expression experiments.
IS50 transposase expression and Tn5 transposition are up-regulated by over-expression of the sRNA SgrS
SgrS down-regulates the expression of several known targets, including the primary glucose transporter encoded by the ptsG gene, a mannose transporter encoded by manXYZ and it up-regulates the expression of yigL, a phosphatase involved in phospho-sugar detoxification . As we observed up-regulation of IS50 transposase expression in cells over-expressing SgrS in M9 glucose media, we considered the possibility that this effect was a response to glucose limitation. In fact, we show in Additional file 2 that induction of SgrS in M9 glucose resulted in a substantial slowing of bacterial growth, as would be expected if nutrients had become growth-rate limiting. To further test the glucose limitation hypothesis, we performed a similar experiment in rich media (LB) and in M9 glucose supplemented with glycerol, a carbon source whose import is not dependent on glucose transporters . We also tested the response of the reporter to over-expression of an SgrS mutant, sgrS1, that is incapable of down-regulating glucose import . In these experiments we used a Tn5 TCF as a reporter in the DBH107 strain background; DBH107 has a complete deletion of the lac operon and consequently the plasmid-encoded sRNA genes are constitutively expressed. To avoid problems in growing these cells, cultures were initially propagated in either LB or M9 glucose/glycerol and then where indicated, switched to other media.
We show in Figure 8B that after approximately 4 hours of SgrS over-expression in M9 glucose, reporter expression increased close to 5-fold relative to a ‘vector’ control. In contrast, over-expression of SgrS1 was incapable of up-regulating reporter expression under these same conditions, suggesting that SgrS must be able to down-regulate glucose import and or retention in order to increase transposase transcription. When cells were grown in M9 glucose supplemented with glycerol, expression of SgrS as above caused only an approximately 2-fold increase in transposase expression. Importantly, the reduced effects of SgrS on transposase expression under ‘glycerol’ conditions cannot be explained by differential expression of the respective sRNAs, as levels of SgrS and SgrS1 were similar in M9 glucose with or without glycerol (Figure 8C). Also, we failed to see significant transposase induction when SgrS was over-expressed in LB media where there are multiple carbon sources. Finally, consistent with the glucose limitation hypothesis, we also show in Figure 8B that increased transposase expression resulting from SgrS expression in M9 glucose was the only condition that inhibited cell growth.
Hfq is a global regulator of gene expression in bacteria. However, until recently, Hfq had not been linked to the control of transposable elements. Work in the Tn10/IS10 system provided the first example of Hfq inhibiting a transposon . In the current work, we asked if the transposition of a related element, Tn5/IS50, is also regulated by Hfq. We show that Tn5 transposition and IS50 transposase expression are repressed by Hfq; however, the mechanism of repression is atypical for Hfq, involving predominantly a block in IS50 transposase transcription. Preliminary evidence is presented that is consistent with Hfq modulating IS50 transposase transcription through regulation of Crp. We also show that transposase transcription and Tn5 transposition are activated by over-expression of the sRNA SgrS and provide evidence that this is a transcriptional response to glucose limitation.
Hfq negatively regulates Tn5 transposition
The results of ‘mating out’ experiments were consistent with Hfq acting as a strong negative regulator of Tn5 transposition. Tn5 transposition increased close to 75-fold in one hfq − strain (DBH179 background). The magnitude of this increase was somewhat surprising given that up-regulation of Tn10 in hfq − , under essentially antisense-minus conditions, was about 7-fold . However, in a different hfq − strain (DBH261 background) Tn5 transposition increased only 9-fold. At this point it is unclear why there was such a large discrepancy in the ‘mating out’ values for the two strains. One possibility is that colony counts in the DBH179 ‘mating out’ (hfq − ) included clones that had ‘jack-pot’ events. That is, colonies were counted that did not derive from independent transposition events. This could explain the high standard error associated with the transposition frequency in the hfq − column in Figure 2A. If, for example, we removed the 3 most prominent outliers from the (DBH179) hfq − data set, the fold increase in transposition dropped to 15-fold, which is more in line with what we observed in the DBH261 strain background and for Tn10 in single copy .
A trans-complementation (Figure 2A) experiment provided definitive proof that the increase in Tn5 transposition detected in one of our hfq − ‘mating out’ strains (DBH179 background) was in fact due to hfq deficiency. In addition, the failure of two Hfq RNA-binding face mutants to provide complementation was consistent with Hfq-directed inhibition of Tn5 transposition relying on functions of Hfq required in canonical Hfq-directed regulatory pathways . That is, Hfq must retain the ability to bind both mRNAs and sRNAs to influence Tn5 transposition.
Hfq, Crp and IS50 transposase gene expression
Evidence that hfq status influences IS50 transposase expression came from two types of experiments. First, the expression of transposase-lacZ reporter genes in both transcriptional and translational fusion constructs increased significantly under conditions of hfq deficiency. Second, the steady-state level of the native transposase transcript also increased significantly in hfq − . Importantly, the large increase in steady-state transcript level (11-fold) coincided with a less substantial increase in transposase mRNA stability (less than 2-fold increase in half-life). In addition, up-regulation of reporter expression in hfq − for a TLF was almost completely abrogated when the IS50 transposase promoter was replaced by a heterologous promoter. Taken together, these results are consistent with Hfq (or a factor regulated by Hfq) suppressing IS50 transposase expression predominantly at the level of transcription. Notably the suppressive effect of Hfq on IS50 transposase transcription was remarkably specific, as the level of a second transcript (T2) encoded by IS50 was not affected by hfq status.
As Hfq does not typically act directly in gene transcription, we think it likely that Hfq acts indirectly on the IS50 transposase promoter. In addition to DAM, only one other transcription factor, LexA, has been implicated as a regulator of transposase transcription. There is a weak LexA-binding site in the transposase promoter (Figure 1A); however, lexA deficiency was shown to increase transposase transcription only two to three-fold in a TCF . As we have seen increases in transposase expression of up to 11-fold for a TCF in hfq−, it seems unlikely that Hfq would be working through LexA. In contrast, transposase expression increased in dam − to a level more in line with that observed in hfq − (less than two-fold difference in the TCF). However, the observed synergy between hfq − and mutations that rendered the IS50 transposase promoter DAM-insensitive led us to conclude that Hfq does not regulate IS50 transcription by impacting DAM levels (and, therefore, promoter methylation). These results provided motivation to search for other targets of Hfq that impinge on IS50 transposase transcription. This search identified Crp as an additional negative regulator of IS50 transposase transcription. Notably, transposase expression increased to approximately the same level in crp − and hfq − in the experiment in Figure 7. The similar magnitude of up-regulation of transposase expression in hfq− and crp− could be indicative of Hfq acting upstream of Crp to inhibit transposase expression. We did in fact find evidence of Hfq positively regulating Crp protein levels (Figure 7C). This observation is consistent with work recently published in the Y. pestis system where it was found that Crp protein levels decreased approximately five-fold in an hfq disruption strain .
Crp is a known activator/repressor of transcription  and, therefore, more likely than Hfq to be directly involved in regulating IS50 transposase expression at the transcriptional level. Given our evidence that Hfq positively regulates crp expression, a plausible scenario explaining our expression data is that the observed up-regulation of IS50 transposase transcription in hfq − is a result of decreased Crp protein levels. Crp may act either directly or indirectly on the IS50 transposase promoter to repress transcription. This is currently a working model as we have not yet tested the possibility that Crp binds the IS50 transposase promoter and it may only be coincidental that transposase expression increased to similar levels in hfq − and crp − strains. Notably, we also found that Tn5 transposition increased when the crp gene was disrupted, although the extent of the increase was smaller than that observed in the isogenic hfq disruption strain. This could be indicative of additional factors in the Hfq regulon impinging on Tn5 transposition.
There is precedent for Crp down-regulating the transcription of a transposase gene. In the case of IS2, transposase transcription increased close to 200-fold in crp − . It was also shown through protein-DNA footprinting that Crp binds directly to the IS2 transposase promoter . Interestingly, based on the consensus binding sequence for Crp, the authors of the above study predicted that Crp would bind to the IS50 transposase gene. However, the predicted crp binding site is located downstream of the transposase promoter and is not present in our TCF (where we detected increased transposase expression in crp − ). Nevertheless, it would be worthwhile to test for Crp binding to the IS50 transposase promoter as the results of Crp ChIP-chip studies revealed the presence of thousands of weak crp binding sites scattered throughout the E. coli genome . It is also possible that Crp acts indirectly on the IS50 transposase promoter by regulating the expression of another transcription factor.
Tn5 transposition and metabolic stress
We also identified conditions that activate transposase expression and transposition; over-expression of the sRNA SgrS increased transposase expression and transposition approximately five-fold. We favor the possibility that this induction is a consequence of glucose limitation but cannot formally rule out the possibility that SgrS targets an as yet undefined regulatory pathway that impinges on transposase expression. Our reasoning for this is that we observed induction of transposase expression and transposition specifically when cells were grown with glucose as the major carbon source and SgrS is known to prevent expression and function of the major glucose transporter encoded by the ptsG gene . Consistent with this idea, we found that transposase induction levels correlated with a reduced growth rate. Furthermore, we demonstrated that: (i) an allele of SgrS (sgrS1) that is incapable of down-regulating ptsG expression failed to induce transposase expression in M9 glucose; (ii) under conditions where SgrS was expressed in M9 glucose media supplemented with glycerol, we failed to see induction of transposase expression to the same extent as when glycerol was absent; (iii) SgrS expression did not impact transposase expression when cells were grown in rich media (LB) and (iv) over-expression of 3 other sRNAs (RybB, RyeB and MicC) that are not expected to influence glucose transport did not increase transposase expression in M9 glucose -. Precedent for nutritional stress influencing transposition comes from earlier work in the IS903 system where mutations in a gene (aspA) required for fermentative metabolism during anaerobic growth caused transposition to occur at an accelerated rate .
At this point it is unclear as to what factors are driving the induction of the transposase gene under SgrS over-expression conditions. With regard to further defining the mechanism of IS50 transposase up-regulation under SgrS over-expression conditions, it would also be advantageous to find alternative experimental conditions for achieving this increased expression. If, for example, simply starving cells by restricting a carbon source during growth achieves the same end as over-expressing SgrS in M9 glucose media, an unbiased screen to search for genetic factors that are necessary for the up-regulation of transposase expression could be performed to reveal the regulatory network impinging on the transposase promoter. As it stands, any factors that influence SgrS expression would interfere with the outcome of such a screen. Alternatively, if it was found that restricting glucose is not sufficient for inducing transposase expression, the possibility that SgrS plays a more direct role in controlling transposase expression would have to be considered.
In this work, we have identified several genes that impact on IS50 transposase expression, including hfq, crp and sgrS. Hfq and Crp proteins are negative regulators and SgrS RNA (under specific growth conditions) is a positive regulator of transposase gene expression. Exactly how these factors impinge on transposase expression remains to be worked out and at this point it is not clear if we are seeing modulation of the same regulatory network in opposite directions when hfq and crp genes are disrupted and SgrS RNA is over-expressed. Tn5/IS50 is the second transposon identified that is affected by disruption of the hfq gene and the first that does not encode an antisense RNA. This raises the possibility that Hfq influences the transposition frequency of many other bacterial transposons.
Plasmids, bacteriophage and strains
The IS50 translational fusion plasmid (pDH798) is a pWKS30-derivative containing base pairs 1 to 431 of IS50 (nucleotides 1 to 366 of T1) fused to codon 10  of the E. coli lacZ gene. The IS50 transcriptional fusion plasmid (pDH682) is a pUC18-derivative containing base pairs 1 to 80 of IS50 (nucleotides 1 to 15 of T1) fused to nucleotide -16 (relative to the translational start codon) of lacZ. Plasmids encoding sRNAs (pDH764, sgrS; pDH766, rybB; pDH768, micC; pDH772, ryeB) and the corresponding empty vector control (pDH763) were kindly provided by S Gottesman. The plasmid encoding sgrS1 (pDH895) was kindly provided by C Vanderpool. Plasmids encoding Hfq (pDH700, wt) and mutant derivatives (pDH701, K56A; pDH713, Y25A) are described in Ross et al. Details of plasmid constructions are provided in Additional file 3 and a list of oligonucleotides used in this work is provided in Additional file 4.
Lambda phages encoding IS50 transcriptional (λDBH849 and λDBH888) and translational (λDBH812) reporters were generated by cloning IS50 expression cassettes marked with an antibiotic resistance gene (either kanR or cmR) into the his operon of pNK81 and then infecting a strain harboring one of these plasmids with λNK1039, which also contains the his operon. Antibiotic resistant lysogens from the above crosses were selected by replica plating and subsequently phage released from the lysogens were purified, giving rise to λDBH849 (IS50-lacZ-kanR TCF), λDBH888 (IS50-lacZ-CmR TCF) and λDBH812 (IS50-lacZ-KanR TLF).
Plasmids, bacteriophage and strains
Strain or Plasmid
Source or reference
HB101 [F−leu−pro−]; StrR
‘Mating out’ recipient
Source of zjc::Tn5
NK5830 [recA−arg−/F’ lacpro+] zjc::Tn5; KanR
‘Mating out’ donor
DBH179 hfq-1::Ωcat; KanRCmR
‘Mating out’ donor
Source of pOX38Gen
HW-5 [phoA4(Am) his-45 recA1 rpsL99 met-54 F−]; StrR
DBH259 zjc::Tn5; StrRGenRKanR
‘Mating out’ donor
DBH261 hfq-1::Ωcat; StrRGenRKanRCmR
‘Mating out’ donor
DBH261 dam::Tn9cat; StrRGenRKanRCmR
‘Mating out’ donor
MC4100 [F− Δ(argF-lac)169* rpsL150]; StrR
DBH265 hfq-1::Ωcat; StrRCmRKanR
DBH265 dam::Tn9cat; StrRCmRKanR
DBH281 hfq-1::Ωcat; StrRCmRKanR
DBH281 dam::Tn9cat; StrRCmRKanR
DBH303 Δhfq 722::kan; StrRCmRKanR
DBH303 Δcrp 765::kan; StrRCmRKanR
DBH303 Δlrp 787::kan; StrRCmRKanR
NK5830 [recA−arg−/F’ lacpro+]
DBH33 hfq-1::Ωcat; CmR
DBH33 dam::Tn9cat; CmR
DBH238 hfq-1::Ωcat; KanRCmR
DBH238 dam::Tn9cat; KanRCmR
DBH208 hfq-1::Ωcat; KanRCmR
DBH208 dam::Tn9cat; KanRCmR
DBH107 recA−; StrR
DBH107 recA−hfq-1::Ωcat; StrRCmR
DBH33 Δcrp 765::kan
DBH242 Δcrp 765; KanS
DBH344 zjc::Tn5; Kan R
‘Mating out’ donor
W3110mlc rne-Flag-cat; rifSCmR
RNA half-life measurements
W3110mlc rne-Flag-cat Δhfq; rifSCmR
RNA half-life measurements
pSC101-derived; low copy-number ori ; ApR
‘Empty vector’ for Hfq expression
pWKS30-P3-hfq WT ; ApR
pWKS30-P3-hfq K56A ; ApR
pWKS30-P3-hfq Y25A ; ApR
pUC18-derivative; Tn5 t’ase M56A; ApRCmR
Source of Tn5 transposase (No Inh.)
pDH533 with t’ase mutated to G53A,C61A; ApRCmR
pDH533 with t’ase mutated to D97A; ApRCmR
pBR333-derivative; encodes his operon; ApR
pUC18-derivative; IS50-lacZ TCF; ApR
Source of TCF
pDH682-derivative; TCF ‘marked’ with kanR
Parent of pDH849
pDH682-derivative; TCF ‘marked’ with cmR
Parent of pDH888
TCF-kanR from pDH682 cloned into BclI-cut pNK81; ApRKanR
For crossing TCF onto λ
TCF-cmR cloned onto BclI-cut pNK81; ApRCmR
For crossing TCF onto λ
pRZ9905-derivative; full-length IS50-lacZ TLF; ApR
Parent of pDH795
pDH658-derivative; ‘deletion’ TLF used in this study; ApR
Parent of pDH804
pDH795-derivative; TLF ‘marked’ with kanR
Parent of pDH812
TLF-kanR cloned into BclI-cut pNK81; ApRKanR
For crossing TLF onto λ
pWKS30-derivative; contains IS50-lacZ TLF from pDH658; ApR
Parent of pDH798
Vector for sRNA-induction
Encodes his operon
IS50-lacZ translational fusion (TLF) from pDH812 marked with kanR
Chromosomal TLF construction
IS50-lacZ transcriptional fusion (TCF) marked with kanR
Chromosomal TCF construction
IS50-lacZ transcriptional fusion (TCF) marked with cmR
Chromosomal TCF construction
‘Mating out’ assay
Conjugal ‘mating out’ experiments were performed essentially as described for single-copy chromosomal transposons in Ross et al. , except that for measuring transposition in hfq − versus wt, donor growth was carried out in M9 glucose media supplemented with kanamycin (25 μg/mL) and amino acids, instead of LB. DBH13 was used as the recipient. Total exconjugants and transposition events with DBH179 and derivatives were scored by plating mating mixes on M9 glucose plates supplemented with leucine, thiamine and streptomycin (150 μg/mL) or streptomycin and kanamycin (25 μg/mL), respectively. Total exconjugants and transposition events with DBH261 and derivatives were scored by plating mating mixes on M9 glucose plates supplemented with leucine, thiamine, streptomycin (150 μg/mL) and gentamicin (12.5 μg/mL) or streptomycin, gentamicin and kanamycin (25 μg/mL), respectively.
Cells were grown in M9 glucose (with arginine and thiamine) or LB. In situations where strains contained plasmids, plasmids were maintained by including the appropriate antibiotic. Overnight cultures (0.05 mL) were used to seed subcultures (1.5 mL), which typically were grown to mid-log phase before being processed for the Miller assay as previously described .
RNA isolation, primer extension and Northern blot analysis
Total RNA was isolated essentially as described in . For steady-state analysis, cells were grown to mid-log phase in LB before RNA isolation. For half-life analysis, rifampicin (dissolved in dimethyl sulfoxide; DMSO) was added to cell cultures (to 200 μg/mL) to arrest transcription and RNA was isolated immediately before and after rifampicin addition at the indicated time intervals. Primer extension analysis was carried out using 32P-labeled primers oDH230 and oDH390, end-labeled with OptiKinase (USB, Cleveland, OH, USA) according to manufacturer’s instructions. Extension reactions used 5 μg of RNA, and Superscript III reverse transcriptase essentially as described in , except that annealing was performed at 65°C (with no ice treatment) before extending at 55°C for 45 minutes. Extension products were resolved on 6% and 10% denaturing polyacrylamide gels. For Northern blot analysis, 2 μg of RNA was mixed with an equal volume of denaturing load dye (95% deionized formamide [v/v], 10 mM EDTA, 0.5× TBE, 3% xylene cyanol [w/v]), heated to 95°C for 2 minutes, and resolved on a 6% polyacrylamide gel containing 7 M urea. Separated RNAs were electro-transferred to Hybond N (GE Healthcare, Mississauga, ON, Canada) in 0.5× TBE and fixed with UV. Annealing and washing was performed in ULTRAhyb buffer (Ambion, Burlington, ON, Canada) according to the manufacturer’s instructions, using RNA probes complimentary to SgrS or the 5S rRNA (internal standard). To construct the radiolabeled RNA probes, DNA templates for in vitro transcription were made by PCR with primers oDH232/233 (SgrS) and oDH234/235 (5S rRNA) - note that, for each primer pair, the forward primer includes the T7 core promoter. These templates were transcribed in vitro in the presence of 32P-UTP to generate uniformly labeled RNA probes. In vitro transcription reactions were performed in 25 μL volumes with approximately 1 μg DNA template, 1 × T7 RNA polymerase buffer (NEB, Beverly, MA, USA), 20 units RNasin (Promega, Madison, WI, USA), 4 mM dithiothreitol (DTT), 0.16 mg/mL BSA, 0.4 mM each of GTP, CTP and ATP, 0.01 mM UTP, 50 μCi [α-32P]UTP, and 100 units of T7 RNA polymerase.
Cells were centrifuged (2 minutes at 21,000 × g), resuspended in SDS load mix (2% [w/v] SDS, 10% [v/v] glycerol, 50 mM Tris-HCl pH 6.8, 0.25% [w/v] bromophenol blue, 0.8 M β-mercaptoethanol) and heated at 95°C for 5 minutes. To normalize for differences in growth between the various samples, the OD600 of each sample was measured and the volume spun normalized to give an equivalent to OD600 approximately equal to 0.35. The resulting lysates were subjected to SDS-PAGE on a 12% polyacrylamide gel, proteins transferred to PVDF (Roche, Indianapolis, IN, USA) and Crp was detected by Western blot with a polyclonal rabbit anti-Crp antibody (kind gift of H Aiba). The primary antibody was diluted 1:20,000 in TBST; the secondary antibody (anti-rabbit IgG-horseradish peroxidase (HRP) conjugate; Promega, Madison, WI, USA) was used at 1:5,000. Crp was visualized with a Pierce ECL 2 Western blotting substrate (Thermo Scientific, Rockford, IL, USA) and PhosphorImager (GE Healthcare). The membranes were stripped and GroES detected (rabbit anti-GroES antibody from Sigma-Aldrich (St Louis, MO, USA) at 1:10,000) for use as an internal standard; GroES is not sensitive to hfq status . Bands were quantified using ImageQuant software (GE Healthcare) and Crp levels plotted relative to GroES.
bovine serum albumin
cyclic AMP-receptor protein
DNA adenine methylase
polyacrylamide gel electrophoresis
polymerase chain reaction
reverse transcription polymerase chain reaction
sodium dodecyl sulfate
Tris borate-EDTA buffer
20 mM Tris-HCl (pH 7.5), 150 mM sodium chloride, 0.5% Tween-20
translation initiation region
We thank Michael Ellis for providing comments on the manuscript and for useful discussions, and Claire Young for assistance with P1 transductions. We also thank S Gottesman and C Vanderpool for providing pLlacO-sRNA expression plasmids and the sgrS1 allele, respectively, and H Aiba for providing an anti-Crp antibody. Finally, we thank W Reznikoff for providing IS50 transposase plasmids and T Naas for providing an E. coli strain harboring pOX38-Gen. This work was supported by a grant to DBH (MOP 11281) from Canadian Institutes of Health Research. JR and CM were supported by OGS and NSERC scholarships.
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