bdm

Escherichia coli ribonuclease III activity is downregulated byosmotic stress: consequences for the degradation of bdmmRNA in biofilm formationmmi_6986 413..425

Se-Hoon Sim,1† Ji-Hyun Yeom,1† Choy Shin,2

Woo-Seok Song,1 Eunkyoung Shin,1 Hong-Man Kim,1

Chang-Jun Cha,3 Seung Hyun Han,4 Nam-Chul Ha,5

Si Wouk Kim,6 Yoonsoo Hahn,1 Jeehyeon Bae7 andKangseok Lee1*1Department of Life Science (BK21 program) and2Graduate School of Education, Chung-Ang University,Seoul 156-756, Republic of Korea.3Department of Biotechnology, Chung-Ang University,Anseong 456-756, Republic of Korea.4Department of Oral Microbiology and Immunology,Dental Research Institute, and BK21 program, Schoolof Dentistry, Seoul National University, Seoul 110-749,Republic of Korea.5College of Pharmacy and Research Institute for DrugDevelopment, Pusan National University, Busan609-735, Republic of Korea.6Department of Biomaterials Engineering, BK21 Teamfor Biohydrogen Production, Chosun University,Gwangju, 501-759, Republic of Korea.7Graduate School of Life Science and Biotechnology,CHA University, Seongnam 463-836, Republic of Korea.

Summary

During the course of experiments aimed at identifyinggenes with ribonuclease III (RNase III)-dependentexpression in Escherichia coli, we found that steadystate levels of bdm mRNA were dependent on cellularconcentrations of RNase III. The half-lives of adventi-tiously overexpressed bdm mRNA and the activitiesof a transcriptional bdm‘–’cat fusion were observed tobe dependent on cellular concentrations of RNase III,indicating the existence of cis-acting elements in bdmmRNA responsive to RNase III. In vitro and in vivocleavage analyses of bdm mRNA identified twoRNase III cleavage motifs, one in the 5�-untranslatedregion and the other in the coding region of bdmmRNA, and indicated that RNase III cleavages in the

coding region constitute a rate-determining step forbdm mRNA degradation. We also discovered thatdownregulation of the ribonucleolytic activity ofRNase III is required for the sustained elevation ofRcsB-induced bdm mRNA levels during osmoticstress and that cells overexpressing bdm form bio-films more efficiently. These findings indicate that theRcs signalling system has an additional regulatorypathway that functions to modulate bdm expressionand consequently, adapt E. coli cells to osmoticstress.

Introduction

Bacterial mRNA has been recognized for more than halfa century as an inherently unstable molecule in vivo(Brenner et al., 1961; Gros et al., 1961). This intrinsicproperty of bacterial mRNA is believed to contribute to therapid regulation of gene expression that is required forcells to adapt to environmental changes. Although bacte-rial mRNA is normally unstable, the longevity of mRNAvaries from seconds to hours depending on the RNAmolecules and the physiological state of the cell (Nichol-son, 1999; Bernstein et al., 2004). Many cis- and trans-acting elements that govern the differential stability ofmRNA have been identified. Among them, ribonucleasesplay a pivotal role in modulating mRNA stability. InEscherichia coli, the RNase E (Rne) endoribonuclease isa major ribonuclease that catalyses the initial rate-determining cleavage of a large number of transcripts(Hagege and Cohen, 1997). A recent study has alsoshown that the removal of phosphates from the 5′ end ofmRNA by the pyrophosphatase RppH greatly influencesthe stability of a few hundred of mRNAs (Deana et al.,2008).

In recent years, the ribonuclease III (RNase III) family ofenzymes has emerged as one of the most importantendoribonucleases in the control of mRNA stability inhigher organisms (Lee et al., 2006; Jaskiewicz and Filip-owicz, 2008; Ramachandran and Chen, 2008). Althoughthe RNase III family is divided into three classes, allRNase III family members contain a characteristic ribonu-clease domain commonly called the RNase III domain.

Accepted 16 November, 2009. *For correspondence. [email protected]; Tel. (+82) 2 820 5241; Fax (+82) 2 822 5241.†These authors contributed equally to this work.

Molecular Microbiology (2010) 75(2), 413–425 � doi:10.1111/j.1365-2958.2009.06986.xFirst published online 11 December 2009

© 2009 The AuthorsJournal compilation © 2009 Blackwell Publishing Ltd

Class 1 proteins are E. coli RNase III-like proteins, whichusually contain a single ribonuclease domain and adsRNA-binding domain (dsRBD). Class 2 proteins have adsRBD and two ribonuclease domains that are commonlyreferred to as RNase IIIa and RNase IIIb. Class 3 proteinsare the largest and typically contain two ribonucleasedomains: a dsRBD and an N-terminal DExD/H-box heli-case domain followed by a small domain of unknownfunction (DUF283) and a PAZ domain.

The E. coli RNase III was the first specific double-stranded RNA endoribonuclease discovered (Robertsonet al., 1968). This enzyme is an Mg2+-dependent nucleasethat cleaves phosphodiester bonds, creating 5′-phosphate and 3′-hydroxyl termini with an overhang oftwo nucleotides (Court, 1993). RNase III is best known forits role in ribosomal RNA (rRNA) processing in bacteria(Bram et al., 1980), although it has also been shown toparticipate in initiating the decay of several mRNA speciessuch as rnc mRNA (Bardwell et al., 1989; Matsunagaet al., 1996a) and pnp mRNA (Regnier and Portier, 1986).

In this study we set out to investigate the functional roleof RNase III in mRNA stability, and identified 100 geneswith steady state levels of mRNA that were decreased atleast twofold by increased cellular concentrations ofRNase III. Among these genes, the steady state levels ofbdm (biofilm-dependent modulation) mRNA fluctuatedgreatly in a manner that was independent of the RcsCBDHis-Asp phosphorelay system (Rcs system) (Gottesmanet al., 1985), which has been shown to regulate bdmexpression under osmotic stress (Francez-Charlot et al.,2005; Shabala et al., 2009). Based on our experimentalresults, we demonstrate the existence of an additionalregulatory pathway for bdm expression that is mediatedby RNase III, and that the downregulation of RNase IIIactivity is required for a sustained elevation of bdm mRNAlevels in cells under high osmotic stress.

Results

Identification of genes whose mRNA abundance isdependent on RNase III

The functional and evolutionary conservation of RNaseIII-related enzymes in bacteria and higher organismsattests to their biological importance in RNA processingand degradation. The importance of these enzymes in theregulation of gene expression in higher organisms hasbeen vigorously investigated in recent years. However,the functional role of RNase III-related enzymes in theregulation of gene expression in bacteria has not beensystematically studied. This lack of study is likely due tothe active role of RNase III-related enzymes on the pro-cessing of structural RNAs such as rRNA and tRNA, aswell as the absence of genetic systems that can identify

RNase III target mRNA species without affecting normalcellular growth. For this reason, we wished to establishthe functional role of RNase III in regulating gene expres-sion in E. coli through the use of microarray analyses thatwould identify genes with mRNA levels that were depen-dent on cellular concentrations of RNase III. However, forthis purpose, we could not apply a simple procedure thatutilizes total RNA samples prepared from paired culturesof wild-type and mutant (rnc-deleted in this case) cellsbecause E. coli cells with RNase III gene (rnc) deletedgrow at significantly lower rates than wild-type cellsand, consequently, a comparison of mRNA abundancebetween these two strains resulted in the alteration ofmRNA abundance of several hundreds of growth rate-related genes, such as those involved in major metabolicpathways, DNA replication, transcription and translation(data not shown), all of which masked the identification ofgenes whose mRNA abundance was directly regulated byRNase III. To circumvent this problem, we constructed anrnc-deleted strain harbouring a cloned copy of rnc underan arabinose-inducible promoter (pRNC1). Using thisstrain we determined the cellular concentrations of RNaseIII that did not interfere with normal cellular growth. Theresults showed that normal cellular growth was main-tained when RNase III was expressed in the range of0.1–10.0 times the endogenous level of RNase III found inwild-type cells (Fig. 1). It is not unusual for E. coli cells tomaintain the normal cellular growth in rich media when abroad range of ribonuclease concentrations was estab-lished by genetic manipulations. For instance, the cellularconcentrations of the essential E. coli protein, RNase E,required for normal cellular growth in rich media havebeen reported to range from approximately 0.1 to 3 timesthe level of endogenous RNase E found in wild-type cells(Jain et al., 2002; Yeom and Lee, 2006; Shin et al., 2008).

Based on the results shown in Fig. 1, we isolated totalRNA from paired cultures of cells that expressed 0.1, 1.0or 10.0 times the endogenous level of RNase III (0.1¥,1.0¥ or 10.0¥) for microarray analysis. This procedureallowed us to differentiate genes whose expression wasaltered by different cellular concentrations of RNase IIIfrom those dependent on different cellular growth rates.The results showed that an increased cellular concentra-tion of RNase III from 0.1¥ to 10.0¥ upregulated 87 genesas well as downregulated 100 genes by more than twofold(Fig. 1C and Table S1). The relative mRNA abundance ofthese genes was changed in a manner dependent oncellular concentrations of RNase III (Fig. 1D). To validatethe microarray data, the abundance of four gene tran-scripts that were greatly upregulated (ompF and ptsA) ordownregulated (bdm and mltD) by increased cellular con-centration of RNase III was measured using quantitativeRT-PCR. The results of RT-PCR were well correlated withmicroarray data (Fig. S1). These results indicate that

414 S-H. Sim et al. �

© 2009 The AuthorsJournal compilation © 2009 Blackwell Publishing Ltd, Molecular Microbiology, 75, 413–425

RNase III, best known for its role in processing structuralRNAs such as rRNA and tRNA in bacteria, is activelyinvolved in the processing and degradation of mRNA, andconsequently, the post-transcriptional regulation of geneexpression in E. coli.

RNase III negatively regulates bdm gene expression

Among the genes with steady state mRNA levels thatwere decreased by increased cellular concentrations ofRNase III, the bdm gene attracted our attention for thefollowing reasons. First, the steady state levels of bdm

mRNA were dramatically changed by the alteration ofRNase III concentrations in the cell (Fig. S1). Second, thebdm gene has been known to be transcriptionally regu-lated by the RcsCBD His-Asp phosphorelay system (Rcssystem) (Gottesman et al., 1985) in which transcriptionalactivation of bdm is directly mediated by an activatorprotein RcsB (Francez-Charlot et al., 2005). However, thesteady state levels of rcsB mRNA were not dependent oncellular concentrations of RNase III (Fig. S1). Therefore,the increased steady state levels of bdm mRNA causedby decreased cellular concentrations of RNase III werenot caused by transcriptional activation of a bdm promotercontaining an RcsB responsive element (Francez-Charlotet al., 2005). Third, unlike other RcsB targets that areupregulated by high osmolarity through the RcsB system,induced expression of bdm was not transient and wassustained, a finding that suggested the existence of anunidentified mechanism for sustaining RcsB-dependent,osmoregulation-induced bdm expression (Francez-Charlot et al., 2005).

To test whether bdm mRNA contains cis-acting ele-ments that respond to cellular RNase III activity, we mea-sured the degree of resistance to chloramphenicol of cellsexpressing a transcriptional bdm�–�cat fusion mRNA inthe presence and the absence of RNase III. The transcrip-tional bdm�–�cat fusion construct expresses mRNA con-taining a 5′-untranslated region (5′-UTR) and the coding

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Fig. 1. Effect of cellular RNase III concentrations on E. coli growthand transcript profile.A. Arabinose concentration-dependent expression of RNase III.Cultures of W3110 cells harbouring pKAN6 and W3110rnc cellsharbouring pKAN6 or pRNC1 were grown to the early log phase(OD600 = 0.1) and different concentrations of arabinose were addedto induce the synthesis of RNase III (0.001% arabinose for cellsharbouring pKAN6 and 0.00100%, 0.00050%, 0.00025%,0.00013%, 0.00006% and 0.00000% arabinose for W3110rnc cellsharbouring pRNC1). Culture samples were harvested atOD600 = 0.6–0.8 to obtain total protein for Western blot analysis ofRNase III. The relative abundance of the RNase III protein bandswas quantified by setting the amount of RNase III protein producedin W3110 cells harbouring pKAN6 as one.B. Growth characteristics of cells expressing different levels ofRNase III. Cells were grown as described above and their growthwas monitored by analysing cell density (absorbance at 600 nm) atthe indicated time intervals.C. The hierarchical clustering of genes with relative mRNA levelsthat changed by 2.0-fold or more. Red shades represent anincrease and green shades represent a decrease in the level of thetranscript in W3110 plus pKAN6 grown in the presence of 0.001%arabinose and W3110rnc plus pRNC1 grown in the presence of0.0002% (1.0¥ RNase III) or 0.001% arabinose (10.0¥ RNase III)compared with those grown in the presence of 0% arabinose.Genes with relative mRNA levels that changed by 2.0-fold or morein W3110 plus pKAN6 grown in the presence of 0.001% arabinosewere eliminated for the clustering. Lane 1, 0.1¥ versus 1.0¥ RNaseIII; lane 2, 0.1¥ versus 10.0¥ RNase III; lane 3, 0% versus 0.001%arabinose.D. Graphic representation of changes in mRNA abundance of thegenes used for the hierarchical clustering in lanes 1 and 2 inFig. 1C.

Regulation of bdm expression by RNase III 415


region of bdm that are fused to the coding region of CAT.The fusion mRNA was constitutively expressed from amutant tryptophan promoter (Lee et al., 2001) in a multi-copy plasmid (pBRS1) in wild-type cells (W3110) or cellsdeleted for rnc gene (HT115), and the degree of resis-tance of the cells to chloramphenicol was measured. Theresults showed a good correlation between the activitiesof the bdm�–�cat fusion and the cellular concentration ofRNase III (Fig. 2A), indicating that RNase III-responsivecis-acting elements are present in bdm mRNA. To confirmthat the enhanced resistance to chloramphenicol of rnc-deleted cells expressing bdm�–�cat fusion mRNA was aresult of increased CAT protein synthesis, we quantified

by Western blot the amount of CAT protein in cells in thepresence and absence of RNase III expression. Cellsexpressing both RNase III and bdm�–�cat fusion mRNAshowed a ~3.9-fold decrease in the amount of CAT proteinwhen compared with rnc-deleted cells that expressedbdm�–�cat fusion mRNA (Fig. 2B). We further showed thatthe change in the abundance of the RNase III-dependentbdm mRNA resulted from an alteration of bdm mRNAhalf-life by measuring the decay rate of bdm mRNA incells adventitiously coexpressing bdm mRNA and 0.0¥,0.1¥, 1.0¥ or 10.0¥ RNase III. Northern blot analysesrevealed a more than threefold increase in the half-life ofthe bdm transcript (240 versus 72 s) in cells expressingno RNase III compared with cells that expressed 1.0¥RNase III (Fig. 2C). The half-life of bdm mRNA in cellsexpressing 0.1¥ and 10.0¥ RNase III was 106 and 50 srespectively. Collectively, these findings suggest that theincreased steady state level of bdm mRNA in cells withdecreased RNase III activity resulted from the stabiliza-tion of bdm transcripts.

Effects of cellular concentrations of RNase E on bdmmRNA abundance

We tested whether cellular levels of a single-strandedRNA specific endoribonuclease RNase E, which is knownto play a major role in mRNA decay in E. coli (Apirion,1975; Lee et al., 2003; Carpousis, 2007), have an effecton bdm mRNA abundance. As the RNase E gene (rne) isessential in E. coli, we utilized E. coli strain KSL2000 (Leeet al., 2002), in which the chromosomal rne gene hasbeen deleted and complemented with a construct thatexpresses RNase E from an rne gene under thearabinose-inducible PBAD promoter in the pBAD-RNEplasmid. In the KSL2000 strain, the synthesis of RNase Eis controlled solely by the concentration of arabinose, andcellular RNase E levels can be conditionally knockeddown to ~10% of endogenous RNase E levels withoutsignificantly affecting normal cellular growth in rich media.Steady state levels of bdm mRNA were measured inKSL2000 cells either in the presence or the absence ofarabinose. The results revealed no significant changes inbdm mRNA abundance in cells depleted for RNase Ecompared with cells that expressed endogenous levels ofRNase E, indicating that RNase E is not actively involvedin the decay pathway of bdm mRNA (Fig. 3).

Identification of RNase III cleavage sites in bdm mRNAin vitro

We were unable to detect distinct decay intermediates ofbdm mRNA in a Northern blot analysis of total RNA pre-pared from the cells that overexpressed bdm mRNA andRNase III (data not shown). However, the fact that decayintermediates of mRNAs were not detectable at the time of

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Fig. 2. Downregulation of bdm expression by RNase III.A. Effects of RNase III on the activities of bdm�–�cat fusion. StrainsW3110 and HT115 (W3110 rnc-14::DTn10) containing plasmidpBRS1 that expresses the bdm�–�cat fusion were grown at 37°C toOD600 = 0.6 and 10–105 cells were spotted on LB-agar containing 0or 30 mg of chloramphenicol per ml.B. Western blot analysis of CAT protein. Total protein was obtainedfrom the cultures used in (A) and the amount of CAT, RNase III andribosomal protein S1 were analysed using Western blot. The samemembrane probed for CAT was cut and probed with polyclonalantibodies to RNase III and S1. The relative abundance of CATprotein bands was quantified by setting the amount of proteinproduced by W3110 containing pBRS1 as one.C. Effects of cellular RNase III concentration on bdm mRNA decay.Strains HT115 harbouring pBDM3 and pKAN6 or pRNC1 weregrown at 37°C to OD600 = 0.2 and transcription of bdm and rnc fromthe plasmids were induced by the addition of 0.1 mM IPTG for bdmand 0%, 0.0002%, and 0.001% arabinose for 0.1¥, 1.0¥, and 10.0¥expression of RNase III, respectively, for 2 h. Total RNA sampleswere prepared from the cultures 0, 1, 2, 4 and 8 min after theaddition of rifampicin (200 mg ml-1) and separated byelectrophoresis on 6% polyacrylamide gel containing 8 M urea. Theabundance of bdm mRNA and M1 RNA, the RNA component ofRNase P (Kole and Altman, 1979; Reed et al., 1982), wasmeasured by Northern blotting with 5′ end-labelled primers. Theabundance of M1 RNA was measured to provide an internalstandard for evaluating the amount of total RNA in each lane.



analysis was not unusual as these mRNA decay interme-diates are usually subject to rapid decay. For this reason,cleavages of bdm mRNA by RNase III were demonstratedbiochemically using in vitro synthesized bdm mRNA tran-scripts and purified RNase III. RNase III cleavage of inter-nally 32P-labelled bdm mRNA in vitro generated three majorcleavage products approximately 160, 60 and 55 nucle-otides in length (bands d, g and h respectively, in Fig. 4A)and several minor cleavage products consisting of approxi-mately 260-, 220-, 140-, 120-, 25- and 15-nucleotide-longRNA fragments (bands a–c, e, f, i and j respectively, inFig. 4A). The band a was not well distinguished from theintact substrate in Fig. 4A. However, the band a was clearlyvisible in another autoradiograph prepared from a gel thatwas run for a longer time (Fig. S2A). These results indicatethat several cleavage sites exist in the bdm mRNA tran-script. To identify the cleavage sites, 5′-32P-end-labelledbdm mRNA transcripts were cleaved with RNase III. Thereaction generated one major and three minor cleavageproducts of ~220, ~160, 23 and 14 nucleotides in lengththat corresponded to bands b, d, i and j respectively. Theseresults showed that these RNase III cleavage productsgenerated from the 5′ end-labelled bdm mRNA transcriptscontained the intact 5′-terminus, and that at least fourRNase III cleavage sites existed in bdm mRNA. The RNaseIII cleavage sites that produced the bands i and j wereidentified using an RNA ladder generated by alkalinehydrolysis of 5′ end-labelled bdm mRNA transcripts (firstlane in Fig. 4B). These sites were designated as cleavagesites 1 and 2, and were positioned in the double-strandedregion in the 5′-UTR of bdm mRNA, a finding that wasfurther confirmed by primer extension analysis using invitro cleavage products purified from RNase III cleavagereactions and the 5′ end-labelled primer (+20R) (Fig. 4C).

To identify other cleavage sites, primer extension analyseswere performed as described above with the 5′ end-labelled primers (bdmR and +135R) shown in Fig. 4G. Theresults showed five additional cleavage sites, four of whichwere clustered in a close proximity and designated ascleavage site 4 (I–IV) (Fig. 4D, E and G). Cleavage site 3was positioned in the strand opposite of cleavage site 4 inthe double-stranded region of bdm mRNA. The identifiedcleavage sites were consistent with the size of the cleav-age products detected in Fig. 4A, B and H. As shown inFig. 4G, RNase III cleaved the double-stranded regions inthe secondary structure of bdm mRNA, which wasdeduced using the M-fold program (http://mfold.bioinfo.rpi.edu/cgi-bin/rna-form1.cgi) and was partially confirmed byRNase T1 cleavage (Fig. 4B). Overall, RNase III cleavedbdm mRNA in vitro at specific sites that contained RNase IIIcleavage motifs: one in the 5′-UTR and the other in thecoding region of the mRNA (Fig. 4G). Based on the obser-vation that the cleavage products generated by RNase IIIcleavages in the coding region of bdm (RNA fragments d,g and h in Fig. 4A) rapidly appeared and persisted, weconcluded that these sites (3 and 4) are predominantlyresponsible for determining the rate of bdm mRNA degra-dation in vitro.

To obtain a biological relevance of these RNase IIIcleavage sites identified in the synthetic bdm mRNA by invitro cleavage assays, we performed primer extensionanalyses on bdm mRNA produced in vivo using total RNAsamples prepared from cells adventitiously overexpress-ing bdm mRNA. Extension of the same primers used forprimer extension analyses in Fig. 4C–E produced cDNAproducts that corresponded to the cleavage sites 3 and4-II (Fig. 4F). Other cleavage sites (1, 2, 4-I, 4-III and 4-IV)were not detected (Fig. S2B). The cleavage sites 1 and 2identified in vitro are in a very short helix that would notconform to a canonical cleavage motif of RNase III, whichmight have resulted from either a false structure of bdmmRNA or the intrinsic property of RNase III to tend torandomly cleave long RNA transcripts in vitro (Xiao et al.,2009). The results indicated that at least sites 3 and 4-IIwere actually cleaved by RNase III in vivo, although othersites were not detected either because they were notcleaved by RNase III in vivo, or the decay intermediatesrapidly were degraded by other ribonucleases. Takentogether, analyses of bdm mRNA synthesized in vivo andin vitro imply that sites 3 and 4-II are major RNase IIIcleavage sites that probably determine the rate of bdmmRNA degradation.

To test whether the sites 3 and 4-II are major RNase IIIcleavage sites in vivo, a nucleotide substitution (A to U) atposition 157 of the RNase III cleavage site 4-II in bdmmRNA (bdm-A157U) was introduced in pBDM3 and thehalf-life of bdm and bdm-A157U mRNA and the cleavagespecificity of RNase III were investigated. As shown in

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Fig. 3. Effects of cellular concentration of RNase E on bdmmRNA abundance.A. Western blot analysis of RNase III and RNase E. The samemembrane was probed with anti-RNase E, anti-RNase III andanti-S1 antibody. The S1 protein was used to provide an internalstandard to evaluate the amount of cell extract in each lane.B. RT-PCR analysis of bdm mRNA abundance. An E. coli strainKSL2000 was cultured to OD600 = 0.6 and harvested for total RNAand protein preparation. RNase E in KSL2000 cells were depletedas previously described (Lee et al., 2002). The relative abundanceof bdm mRNA was measured using RT-PCR and shown at thebottom of the gel.



Fig. 5, the half-life of bdm-A157U mRNA was increasedby four times (~1 versus ~4 min) and RNase III was notable to efficiently cleave bdm-A157U mRNA at the cleav-age sites 3 and 4-II. These results demonstrated that thesites 3 and 4-II are major RNase III cleavage sites in vivo.

Osmotic regulation of RNase III activity

Previous studies have shown that, unlike other RcsBtargets that are upregulated by high osmolarity through

the RcsB system, the induced expression of bdm by theRcsB system is not transient (Francez-Charlot et al.,2005). This observation was the basis for suspecting theexistence of an unidentified mechanism that is responsiblefor the sustained induction of bdm expression by RcsB-dependent osmoregulation. This notion prompted us toexamine whether the maintenance of induced bdm expres-sion is related to cellular RNase III activity. In order tomonitor the ribonucleolytic activity of RNase III in cellsunder high osmolarity, we used an E. coli strain (KSC004)

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Fig. 4. Identification of RNase III cleavage sites in bdm mRNA in vitro and in vivo.A. In vitro cleavage of bdm mRNA. One pmol of internally 32P-labelled bdm mRNA was incubated with 0.5–1 mg of purified RNase III, in thecleavage buffer with (III + Mg2+) or without MgCl2 (III). Samples were withdrawn at the indicated time intervals and separated on an 8%polyacrylamide gel containing 8 M urea. Cleavage products are indicated (a–j).B. Identification of RNase III cleavage sites using 5′ end-labelled bdm mRNA. One pmol of 5′ end-labelled bdm mRNA was cleaved withRNase III and analysed as described above. Size markers were generated by alkaline hydrolysis (H) and RNase T1 digestion (T1) of the 5′end-labelled bdm transcript. Cleavage sites identified in (C)– (E) are indicated (1–4).C–E. Primer extension analysis of the in vitro cleaved bdm mRNA. Cleavage products of bdm mRNA purified from (B) were hybridized with 5′end-labelled primers (bdm +20R, bdm +135R and bdmR) and extended.F. Primer extension analysis of bdm mRNA synthesized in vivo. Total RNA was prepared from W3110 and HT115 that endogenously (0.5 MNaCl) or adventitiously (pBDM3) overexpressed bdm mRNA and were hybridized with 5′ end-labelled primer (bdmR). Synthesized cDNAproducts were analysed in an 8% polyacrylamide gel. Sequencing ladders were produced using the same primer used in cDNA synthesis andPCR DNA encompassing the bdm gene as a template.G. Predicted secondary structure of bdm mRNA. The secondary structure was deduced using the M-fold program and an RNase T1 digestionas shown in (B). RNase III cleavage sites identified in (B)-(F) are shown (1–4). Bold arrows indicate the cleavage sites identified in (F).H. Schematic representation of RNase III cleavage products of bdm mRNA. Based on the results from (A) to (F), RNase III cleavage productsand sites of bdm mRNA are indicated.



containing an RNase III target site in single copy of apnp�–�lacZ reporter gene fusion (Robert-Le Meur andPortier, 1992; Kim et al., 2008). It has been shown thatRNase III cleaves the pnp untranslated leader, and conse-quently that b-galactosidase production from this fusionconstruct is increased approximately threefold in cellscontaining the rnc14 mutation, which abolishes RNase IIIexpression (Takiff et al., 1989; Robert-Le Meur and Portier,1992; Beran and Simons, 2001). The activity of the pnp�–�lacZ reporter gene fusion in the KSC004 strain followingan osmotic shift with 0.5 M NaCl was increased approxi-mately twofold from 2–8 h following osmotic upshift com-pared with cells that were not treated with NaCl (Fig. 6A).This suggests that exposure of cells to high osmolarityresulted in downregulation of cellular RNase III activity,which, in turn, resulted in the decreased cleavage of tran-

scripts containing an RNase III-targeted site in pnp mRNAfused to lacZ. To eliminate a possibility that high osmolarityregulates some other process, such as transcription ortranslation of the pnp�–�lacZ reporter used, we testedosmotic regulation of RNase III activity using anotherE. coli strain (KSC006) containing an single copy of arnc�–�lacZ reporter gene fusion (Matsunaga et al., 1996a;Kim et al., 2008). It has been shown that RNase III cleavesits own transcript, and consequently b-galactosidase pro-duction from this fusion construct is increased approxi-mately ninefold in cells expressing non-functional RNaseIII (Matsunaga et al., 1996a,b). Analogous results wereobtained from this experiment (Fig. 6B), indicating thatRNase III activity is downregulated by osmotic stress.

Decreased cellular RNase III activity was not associ-ated with cellular concentrations of RNase III, as evalu-

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ated quantitatively by measuring cellular levels of RNaseIII protein using antibodies specific to RNase III inWestern blot analysis (Fig. 6C). We further tested therequirement of downregulated RNase III activity for sus-taining high levels of bdm mRNA was not associated withcellular levels of YmdB (Fig. 6C), a recently discoveredprotein inhibitor of RNase III that has been shown to be

upregulated during cold shock (Kim et al., 2008). Cellularlevels of YmdB in KSC004 were similar until 4 h anddecreased at 8 h after osmotic shift. We think that thedecreased RNase III activity observed in cells exposed tohigh NaCl was associated with other factors such asuncharacterized protein regulators of RNase III, whichhave been recently reported to exist (Kim et al., 2008).

We further tested the requirement of downregulatedRNase III activity to sustain high levels of bdm expressionin cells under osmotic stress. For this, we utilized anE. coli strain with a chromosome that contains a transcrip-tional fusion between the bdm region and lacZ (lbdm6).The transcriptional fusion lbdm6 contains all the RNaseIII cleavage sites that were identified in the experimentsdescribed in Fig. 4. The activities of this fusion were moni-tored following an osmotic shift with 0.5 M NaCl in thepresence of cellular concentrations 1¥ and 10¥ RNase III.The fusion was activated at a similar degree upon theosmotic shift as previously reported (Francez-Charlotet al., 2005). However, the activities of the fusion weredecreased by approximately twofold when RNase III was10 times overexpressed (Fig. 6D and E). These resultsshow that the downregulation of RNase III activity isrequired to sustain the high levels of bdm mRNA inducedby the RcsB system in cells during osmotic stress.

Enhancement of biofilm formation by induction ofbdm expression

It has been shown that bdm expression is upregulated incells during osmotic stress (Weber and Jung, 2002;Cheung et al., 2003; Francez-Charlot et al., 2005) andaltered in biofilms (Prigent-Combaret et al., 1999), but itsphysiological relevance has not been characterized. Wetherefore designed experiments to characterize the rela-tionship between bdm expression and biofilm formation.The results showed a good correlation between bdmexpression levels and enhancement of biofilm formation.Biofilm formation was enhanced when overexpression ofbdm was induced adventitiously from a multicopy plasmidor endogenously by osmotic upshift, and was inhibitedwhen bdm expression levels were decreased by RNase IIIoverexpression (Fig. 6F). The degree of the enhancedformation of biofilm was higher when bdm overexpressionwas adventitiously induced compared with that by osmoticshift. Analysis of bdm mRNA abundance showed a 2.7-fold increase in the steady state level of bdm mRNApresent in cells that adventitiously overexpressed bdmthan in cells with an osmotic shift (Fig. 6F), indicating thatthe differences we observed in the degree of enhancedbiofilm formation were probably due to the differentexpression levels of bdm mRNA in these cells. The cor-relation coefficient between the relative amount of bdmmRNA and the amount of biofilm formation was 0.94,

B

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Fig. 5. Effects a nucleotide substitution (A157U) in the RNase IIIcleavage site 4-II on the bdm mRNA half-life and RNase IIIcleavage in vivo.A. Effects of A157U on bdm mRNA decay. Strains W3110harbouring pBDM3 or pBDM3-A157U were grown at 37°C toOD600 = 0.2 and transcription of bdm from the plasmids wasinduced by the addition of 0.1 mM IPTG for 2 h. RNA samples wereprepared and analysed as described in the legend of Fig. 2C.B. Effects of A157U on bdm mRNA cleavage by RNase III in vivo.Total RNA was prepared from the cultures described above andanalysed as described in the legend of Fig. 4F.



C

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Fig. 6. Downregulation of RNase III activity by high osmolarity and effect on biofilm formation.A and B. Osmotic regulation of RNase III activity. E. coli strains KSC004 (A) and KSC006 (B) were grown in M63 supplemented with 0.4%glucose and either shocked at time zero with 0.5 M NaCl or left untreated. Cultures were withdrawn at the indicated time intervals and ab-galactosidase assay was performed.C. Western blot analysis of RNase III and YmdB. Total proteins were prepared from the KSC004 cultures grown as described in Fig. 6A,separated on 8% SDS-PAGE and immunoblotted with anti-RNase III, anti-YmdB and anti-S1 protein. The S1 protein was used to provide aninternal standard to evaluate the amount of cell extract in each lane.D. Effects of RNase III overexpression on bdm expression in cells under osmotic stress. E. coli strain lbdm6 containing a single copy of abdm�–�lacZ reporter gene fusion was grown as described in (A), and a b-galactosidase assay was performed.E. Western blot analysis of RNase III. Total proteins were prepared from the cultures grown as described in Fig. 6D and immunoblotting wasperformed as described above.F. Effects of bdm overexpression on biofilm formation. Strains used were: 1. W3110Dbdm plus pKAN6 and pTrc99A (no NaCl); 2. W3110 pluspKAN6 and pTrc99A (no NaCl); 3. W3110 plus pKAN6 and pTrc99A (0.5 M NaCl); 4. W3110 plus pKAN6 and pBDM3 (no NaCl); 5. W3110plus pRNC1 and pBDM3 (no NaCl). Overexpression of bdm was endogenously (3, W3110 plus pKAN6 and pTrc99A) or adventitiously (4,W3110 plus pKAN6 and pBDM3) induced by an osmotic shift with 0.5 M NaCl or addition of 0.1 mM IPTG respectively. 0.1 mM IPTG and0.1% arabinose were added to all the cultures (1–5). Relative activity of RNase III was estimated based on Fig. 6A–D. The degree of biofilmformation was quantified using the crystal violet staining method as described in the Experimental procedures section. Quantification of bdmmRNA abundance was measured using RT-PCR.



which shows a strong interrelationship between them.These findings suggest that bdm is involved in osmoticstress-induced formation of biofilm.

Discussion

Genome-wide analyses of mRNA abundance at single-gene resolution identified 87 genes that were upregulatedand 100 genes that were downregulated by increasedcellular concentrations of RNase III. These genes areprimarily involved in carbohydrate transport and metabo-lism, energy production and conversion, post-translationalmodification, protein turnover and chaperones, illustratingthe effects of RNase III activity on a broad range of cel-lular physiological processes. Expression levels of theseRNase III-dependent genes might be the consequence ofan indirect effect of cellular RNase III concentration, suchas regulation at the transcriptional level. However, thepresent study and other studies have shown that expres-sion of at least some of these genes results from the directaction of RNase III on mRNA (Aristarkhov et al., 1996;Matsunaga et al., 1996a; Robert-Le Meur and Portier,1992).

In the case of adh expression, adh mRNA is processedin the 5′-UTR by RNase III to yield translationally activemRNA, which becomes stable due to ribosomal protectionfrom ribonucleases. We also found that the mRNA ofsome of upregulated genes is cleaved in the 5′-UTR byRNase III to yield translationally active mRNA (S-H. Sim,E. Shin and K. Lee, in preparation). In contrast to mRNAspecies processed by RNase III, RNase III degrades bdmmRNA by cleaving it in several places, which are consti-tuted by two RNase III cleavage motifs, one in the 5′-UTRand the other in the coding region of the mRNA (Fig. 4).Analysis of cleavage products indicated that RNase IIIcleavage sites in the coding region are likely to be respon-sible for determining the rate of bdm mRNA degradation(Figs 4 and 5). Considering that RNase III is a double-stranded RNA-specific endonuclease and that thesecleavage sites (3 and 4-II) are present in the double-stranded region of the bdm coding region, it seems thatbdm mRNA forms an extensive secondary structuresimilar to the one shown in Fig. 4G as a result of poortranslation, and consequently is subject to degradation byRNase III. A recent report also indicated that mRNAfolding plays a predominant role in mRNA stability andtranslation efficiency (Kudla et al., 2009). Taken together,these observations imply that the abundance of a subsetof mRNA species is directly controlled by RNase III.However, comparative analyses of predicted highlyexpressed genes (Karlin and Mrazek, 2000) and RNaseIII-dependent genes failed to show a correlation betweenprotein expression levels and the degree of mRNA abun-dance that is controlled by RNase III (data not shown).

This may reflect the complexity of the mechanisms thatgovern the modulation of RNase III activity upon each ofthe RNase III-targeted mRNA molecules. Multiple cis-and/or trans-acting elements seem to be involved indetermining which of the RNase III-targeted mRNAspecies is degraded or processed, as indicated in thepresent study.

Among the genes downregulated by RNase III, weinvestigated the physiological importance of bdm down-regulation by RNase III and identified an additional regu-latory pathway mediated by modulation of RNase IIIactivity during the RcsB-dependent adaptation of E. colicells to high osmolarity. This novel pathway involvesmodulation of RNase III activity on degradation of bdmmRNA whose product enhances biofilm formation.Involvement of bdm expression in biofilm formation hasbeen suspected, but not demonstrated until the presentstudy. Homologues of Bdm are found in closely relatedbacterial species of E. coli including Shigella dysenteriae,Salmonella enterica ssp., Enterobacter sp. and Klebsiellapneumonia, thus indicating the existence of a conservedfunction of Bdm protein in these bacterial species.

A recently discovered protein inhibitor of RNase IIIactivity, YmdB, which is upregulated during cold shock(Kim et al., 2008), is not involved in osmoregulation ofRNase III activity (Fig. 6C). We believe that at least oneother unidentified trans-acting factor inhibits RNase IIIactivity on bdm mRNA degradation under conditions ofosmotic stress. This view is supported by the report ofseveral different inhibitors of RNase III (Kim et al., 2008),whose identities are not yet described. Trans-actingfactors that inhibit RNase III activity on bdm mRNA deg-radation can be experimentally identified by a geneticapproach similar to that used by Kim et al. (2008). In thecase of RNase E, protein inhibitors that differentiallymodulate the ribonucleolytic activity have been recentlydiscovered (Lee et al., 2003; Gao et al., 2006). Thepresent study demonstrates the ability of RNase III topromote the decay of bdm mRNA, allowing cells to rapidlyadjust to environmental changes, and imply that RNase IIIribonucleolytic activity is differentially regulated bymolecular mechanisms that involve mRNA-specific cis-and/or trans-acting factors.

Experimental procedures

Strains and Plasmids

E. coli strain HT115 (W3110 rnc-14::DTn10) was obtainedfrom Dr D. L. Court. The N3433rnc strain was constructedfrom N3433 by phage P1-mediated transduction using HT115as a donor strain. W3110Dbdm was constructed by deletingthe open reading frame of bdm in the genomic DNA of W3110using the procedure described by Datsenko and Wanner(2000). PCR primer were 5′-bdm-D (5′- AACCCCTAAATTAG



GTTGCCGATCAAGCATAGCACCTTAGTGTAGGCTGGAGCTGCTTC) and 3′-bdm-D (5′- GCAGACACCATAAATACACAGACACGGAGAATCACTATGCATATGAATATCCTCCTTA),and pKD3 (Datsenko and Wanner, 2000) was used as atemplate.

To construct pRNC1, the coding region of rnc was amplifiedusing two primers, rnc-5′ (5′-AGAATTCATATGAACCCCATCGTAATAA) and rnc-3′ (5′-CTCTAGATCATTCCAGCTCCAGTTTTTTCA), digested with NdeI and XbaI and ligatedinto pKAN6B-IF1 (Yeom et al., 2008). Plasmid pBDM3 wasconstructed by subcloning an NcoI and XbaI fragment encod-ing Bdm that was amplified using the primers bdm-UTR-F(5′-ATCCATGGGCTTTATAAATCTGCGATCC) and bdm-R2(5′-CGTCTAGATTAAAGCGTAGGGTGCTGGCCAC) intopTrc99A. Plasmid pBDM3-A157T was constructed by sub-cloning an NcoI and XbaI fragment encoding Bdm that wasamplified using overlap-extension PCR into pTrc99A. Theprimers used were bdm-UTR-F, bdm-R2 bdm +155R (5′-AGGATGTCGTTATCAGAAGCTTCAAC), and bdm-A157T-F(5′-GTTGAAGCTTCTGATAACGACATCCTCTGTGATATCTACCAGCAAACG).

The pBRS1 plasmid that expresses the bdm�–�cat fusionwas constructed in multiple steps. To construct pBRS1, NcoIand NotI sites were created using the overlap-extension PCRmethod with following primers: bdm-reporter 1 (5′-GCATAGCGGCCGCTTTATAAATCTGCGATCCGTAG), bdm-reporter 2 (5′-GGTATATCCAGTGATTTTTTTCTCAAGCGTAGGGTGCTGGCCACTG), Eno3 (5′-GAGAAAAAAATCACTGGATA) and RMC-MscI (5′-CCTTGTCGCCTTGCGTATAA). Two separate PCR products (PCRs 1 and 2) wereobtained using primers, bdm-reporter 1/bdm-reporter 2 andEno3/RMC-Msc I respectively, and the PCR products PCRs 1and 2 were combined and amplified using two outsideprimers bdm-reporter 1 and RMC-Msc 1. The resulting PCRproduct was digested with NotI and NcoI and cloned into thepCAT924. Plasmid pCAT924 was constructed by self-ligatingthe BamHI fragment from pRNA122 (Lee et al., 1997; 2001).E. coli strains KSC004 (Robert-Le Meur and Portier, 1992;Kim et al., 2008) and KSC006 (Matsunaga et al., 1996a; Kimet al., 2008) were obtained from Dr Stanley N. Cohen. E. colistrain lbdm6 (Francez-Charlot et al., 2005) was obtainedfrom Dr Kaymeuang Cam.

Microarray procedures

Relative mRNA levels were determined by parallel two-colour hybridization to DNA oligonucleotide microarrays onglass slides, which have been produced and provided bythe 21st Century Frontier Microbial Genomics and Applica-tion Center Program of the Korean Ministry of Science(http://www.microbe.re.kr/). Comparisons between pairedcultures that expressed 0.1¥ versus 1.0¥ and 0.1¥ versus10.0¥ RNase III proteins relative to the expression level ofendogenous RNase III were performed directly. To identifygenes whose mRNA abundance was changed due to thedifferent concentrations of arabinose present in the cultures,comparisons between paired cultures of wild-type cellsgrown in the presence of 0% and 0.001% arabinose werealso performed. Synthesis of cDNA and hybridization wereperformed by Digital Genomics (Seoul, Korea). We mea-sured the relative mRNA abundance under appropriate con-

ditions in two replicate comparisons, selecting genes whoserelative mRNA levels were changed twofold or more andfurther clustering them by the previously described method(Eisen et al., 1998).

Western blot analysis

Western blot analysis was carried out as described previously(Kime et al., 2008). Polyclonal antibodies to RNase III wereobtained from rabbits inoculated with purified His-taggedRNase III as described (Amarasinghe et al., 2001). Poly-clonal antibodies to YmdB were obtained from Dr Stanley N.Cohen. Specific proteins were imaged using VersaDoc 100(Bio-Rad) and quantified by Quantity One (Bio-Rad).

RT-PCR

The procedure for RT-PCR analysis has previously beendescribed (Yeom and Lee, 2006; Yeom et al., 2008). Thefollowing primers were used: 5′-ATGTTTACTTATTATCAGGCAG and 5′-TTAAAGCGTAGGGTGCTGGCCAC forbdm, 5′-ATGAAGGCAAAAGCGATATTAC and 5′-TCAGGAATCTGGCATGTTGTTG for mltD, 5′-ATGATGAAGCGCAATATT and 5′-TTAGAACTGGTAAACGAT for ompF,5′-ATGGCCCTGATTGTGGAA and 5′-TTACAGTTCCAGTTCATG for ptsA and 5′-ATGAACAATATGAACGTA and5′-TTAGTCTTTATCTGCCGG for rcsB.

Northern blot analysis

Total RNA samples were prepared from the cultures using anRNeasy mini prep kit (Qiagen) following 0, 2, 4 and 8 minafter addition of rifampicin (200 mg per ml). Twenty micro-grams of the total RNA sample was denatured at 70°C for10 min in an equal volume of formamide loading buffer andseparated by electrophoresis on a 6% polyacrylamide gelcontaining 8 M urea. The procedure for Northern blot analysishas previously been described (Lee et al., 2002). The oligo-nucleotide probes used for probing M1 RNA and bdm mRNAwere M1 (5′-GCTCTCTGTTGCACTGGTCG) and bdm (5′-ATGTTTACTTATTATCAGGCAG) respectively.

In vitro cleavage assay

His-tagged RNase III purification and cleavage assays wereperformed as previously described (Amarasinghe et al.,2001). One pmol of labelled RNA were incubated with0.5–1 mg of purified RNase III in the presence of 0.25 mg ml-1

of yeast tRNA (Ambion) and 20 U of RNaseOUT (Invitrogen)in cleavage buffer (30 mM Tris-HCl, pH 7.9, 160 mM NaCl,0.1 mM DTT, 0.1 mM EDTA pH 8.0). Cleavage reactionswere initiated by adding 10 mM MgCl2 after 5 min of incuba-tion at 37°C. Samples were removed at time intervals andmixed with an equal volume of Gel Loading Buffer II (Ambion)containing 95% of formamide, 18 mM EDTA, 0.025% SDS,0.025% xylene cyanol and 0.025% bromophenol blue. Thesamples were denatured at 65°C for 10 min and separatedon an 8% or 10% polyacrylamide gel containing 8 M urea and1¥ TBE.



Primer extension analysis

In vitro cleaved RNA was purified by phenol extraction andethanol precipitation, and hybridized with 5′-32P-labelledprimers. The following primers were used: bdm +20R (5′-GCCTGATAATAAGTAAACAT), bdm +135R (5′-TTCAACCCATTTGGTCAGCT) and bdm-R (5′-TTAAAGCGTAGGGTGCTGGC). RNA and labelled primers were annealedat 65°C for 10 min then slowly cooled down to 37°C for 1–2 h,and extended at 37°C for 1 h using AMV reverse transcriptase(New England Biolabs, USA). The extended fragments wereseparated on polyacrylamide gels as described above.

b-Galactosidase assay

b-Galactosidase activity in whole cells was determined asdescribed previously (Miller, 1992).

Quantification of biofilm formation

Biofilm formation was measured by the ability of the cells toadhere to the wells of 96-well PVC microtiter plates (BDfalcon) as previously described (Djordjevic et al., 2002).Overnight cultures were diluted in LB medium at OD600 = 0.1,0.1 mM IPTG and/or 0.001% arabinose were added to inducetranscription from plasmid-born promoters and cultured toOD600 = 0.7–0.8. For osmotic upshift, NaCl was added to afinal concentration of 0.5 M. The cultures were diluted 1:10 inminimal M9 medium containing 0.4% glucose, and 0.2 ml ofthe diluted cultures was transferred into PVC microtiter platewells per strain. The plate was incubated at 30°C overnightfor biofilm formation. After a 10 or 20 h incubation period,medium was removed from the well using micropipette.Plates were air-dried for 30 min, 20 ml of 1% crystal violet wasadded into each well. Plates were placed at room tempera-ture for 20 min and each well was rinsed thoroughly withwater followed by an addition of 200 ml of 95% ethanol todestain the wells. The absorbance of the crystal violet stainedbiofilm was measured at 595 nm in a spectrophotometer.

Acknowledgements

We are grateful to Dr Allen W. Nicholson for helpfulcommentary. This work was supported by grants fromNational Research Foundation of Korea Grant funded by theKorean Government (2009-0065181), the 21C FrontierMicrobial Genomics and Application Center Program of theMinistry of Education, Science and Technology, Republic ofKorea, and the Pioneer Research Program for ConvergingTechnology of the Ministry of Education, Science and Tech-nology, Republic of Korea (M1071118001-08M1118-00110).

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