JOURNAL OF BACTERIOLOGY, Mar. 2010, p. 1671–1684 Vol. 192, No. 60021-9193/10/$12.00 doi:10.1128/JB.01311-09Copyright © 2010, American Society for Microbiology. All Rights Reserved.

MreB Drives De Novo Rod Morphogenesis in Caulobacter crescentusvia Remodeling of the Cell Wall�†

Constantin N. Takacs,1‡ Sebastian Poggio,1 Godefroid Charbon,1§ Mathieu Pucheault,1¶Waldemar Vollmer,2 and Christine Jacobs-Wagner1,3,4*

Department of Molecular, Cellular and Developmental Biology, Yale University, New Haven, Connecticut 065201; Centre forBacterial Cell Biology, Institute for Cell and Molecular Biosciences, Newcastle University, Newcastle upon Tyne,

NE2 4HH, United Kingdom2; Section of Microbial Pathogenesis, Yale School of Medicine, New Haven,Connecticut 065203; and The Howard Hughes Medical Institute, New Haven, Connecticut 065204

Received 2 October 2009/Accepted 4 December 2009

MreB, the bacterial actin-like cytoskeleton, is required for the rod morphology of many bacterial species.Disruption of MreB function results in loss of rod morphology and cell rounding. Here, we show that the widelyused MreB inhibitor A22 causes MreB-independent growth inhibition that varies with the drug concentration,culture medium conditions, and bacterial species tested. MP265, an A22 structural analog, is less toxic thanA22 for growth yet equally efficient for disrupting the MreB cytoskeleton. The action of A22 and MP265 isenhanced by basic pH of the culture medium. Using this knowledge and the rapid reversibility of drug action,we examined the restoration of rod shape in lemon-shaped Caulobacter crescentus cells pretreated with MP265or A22 under nontoxic conditions. We found that reversible restoration of MreB function after drug removalcauses extensive morphological changes including a remarkable cell thinning accompanied with elongation,cell branching, and shedding of outer membrane vesicles. We also thoroughly characterized the compositionof C. crescentus peptidoglycan by high-performance liquid chromatography and mass spectrometry and showedthat MreB disruption and recovery of rod shape following restoration of MreB function are accompanied byconsiderable changes in composition. Our results provide insight into MreB function in peptidoglycan remod-eling and rod shape morphogenesis and suggest that MreB promotes the transglycosylase activity of penicillin-binding proteins.

Most bacteria have characteristic cell morphologies main-tained during growth (67). The peptidoglycan (PG) componentof the cell wall represents in most cases the physical support ofvarious bacterial shapes. PG is a mesh-like polymeric macro-molecule which opposes the osmotic pressure of the bacterialcytoplasm and prevents lysis in hypotonic growth environments(29). Isolated PG cell walls (sacculi) retain the shapes of thecells from which they originate while PG disruption causes theformation of osmotically labile spheroplasts, underscoringPG’s essential role in cell shape determination and cellularintegrity maintenance. PG is composed of long glycan chainsthat are oriented roughly along the short axis of rod-shapedGram-negative bacteria and that are connected by short pep-tide cross-links (21, 60). Bacterial growth and division neces-sitate the expansion and division of the PG cell wall, whichrequires the insertion of new PG material in the preexisting,covalently linked mesh (29). New PG synthesis requires two

enzymatic reactions performed by penicillin-binding proteins(PBPs). Glycan chain synthesis is achieved by transglycosyla-tion activity while cross-linkage of glycan chains to the existingmesh is achieved by transpeptidation activity (47). Class APBPs, called bifunctional or bimodular PBPs (e.g., PBP1a and1b of Escherichia coli), possess both transpeptidase and trans-glycosylase domains while class B PBPs, such as PBP2 andPBP3 of E. coli, can perform only transpeptidase reactions(47). Controlled degradation of the PG by cell wall hydrolasesis necessary for incorporation of new PG material duringgrowth. Tight coordination between PG synthesis and degra-dation is required to maintain the integrity of the mesh at alltimes (29).

The bacterial cytoskeleton also plays a central role in cellshape determination and maintenance (7). MreB is a bacterialactin homolog that forms dynamic helical structures under-neath the cytoplasmic membrane in most rod-shaped bacteria(8, 34, 37, 56). In some species, the spatial distribution of MreBvaries during the cell cycle, changing from a helical/patchylocalization pattern throughout the cell to a ring-like distribu-tion near midcell (20, 22, 50, 58). MreB is required for rodshape maintenance as deletion of the MreB-encoding gene ordepletion of MreB causes loss of rod shape and cell rounding(20, 22, 34, 63). Other proteins, including MreC, MreD, RodA,PBP2, and RodZ, function along with MreB to maintain rodshape as loss of their function also results in cell rounding (2,5, 33, 48, 62). Among these rod-morphogenic proteins, onlyPBP2 has a known enzymatic function, being involved in PGsynthesis as an elongation-specific transpeptidase; the others

* Corresponding author. Mailing address: Department of Molecu-lar, Cellular, and Developmental Biology, KBT 1032, Yale University,P.O. Box 208103, New Haven, CT 06520-8103. Phone: (203) 432-5170.Fax: (203) 432-6161. E-mail: christine.jacobs-wagner@yale.edu.

‡ Present address: The Rockefeller University, New York, NY10065.

§ Present address: Department of Molecular and General Physiol-ogy, Roskilde University, DK-4000 Roskilde, Denmark.

¶ Present address: CPM UMR 6510, CNRS, Universite de Rennes1, 35042 Rennes Cedex, France.

† Supplemental material for this article may be found at http://jb.asm.org/.

� Published ahead of print on 18 December 2009.

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are membrane-spanning or integral membrane proteins (2, 5,15, 48). The overall involvement of these morphogenetic pro-teins in rod shape maintenance has led to a model in whichthey are part of the elongase complex, a PG synthesizing ma-chine that elongates the PG side wall (2, 5, 15, 48, 59). Theelongase complex would include PG lytic enzymes and at leastone bifunctional PBP required for glycan strand synthesis (15,59). In Bacillus subtilis, MreB homologs were found to associ-ate with the bifunctional PBP1 (36) and to regulate the local-ization of the PG hydrolase LytE (9). However, it is still un-clear how MreB functions in the context of the proposedelongase complex to determine and maintain rod shape.

It has been previously shown that repletion of MreB inlemon-shaped, MreB-depleted Caulobacter crescentus cellsleads to the formation of cell filaments that present branchesand ectopic stalks (64). To examine how MreB can drive denovo rod shape morphogenesis, we followed a similar strategyexcept that we used drug treatment to interfere with MreBfunction. The small molecule 3,4-dichlorobenzyl carbamimido-thioate, also known as A22, has been shown to rapidly disruptMreB localization in vivo and to induce growth-dependentrounding in several Gram-negative bacteria (23, 32, 41, 45, 52).Furthermore, genetic and biochemical experiments haveshown that MreB is the direct molecular target of A22 and thatA22 binds to MreB’s ATP-binding pocket, inducing a statewith low affinity for polymerization (3, 23). As removal of A22is followed within minutes by recovery of the normal MreBlocalization pattern (23), this drug represents a convenient toolfor rapid and reversible inhibition of MreB function. However,A22 was found to inhibit the growth of an mreB deletionmutant of E. coli, suggesting that it can have MreB-indepen-

dent toxic effects (35). In this study, we show that the toxicityof A22 varies with the drug concentration, culture mediumconditions, and Gram-negative species tested. We identify asimilarly potent but less toxic structural analog, MP265 (4-chlorobenzyl carbamimidothioate), as well as nontoxic concen-trations and conditions for both A22 and MP265 that induceloss of rod cell morphology in C. crescentus. We also show thatrecovery of rod shape after drug removal is accompanied byintensive remodeling of PG morphology and composition.

MATERIALS AND METHODS

Strains, plasmids, and growth conditions. The strains and plasmids used inthis study are listed in Table 1. To construct pVsigpepCHYN-4, the gene regioncoding the signal peptide of CC1996 was PCR amplified from C. crescentusgenomic DNA using primers 5�-CACATATGATGAGGCAGTTGTGGACGCAAG-3� and 5�-CACATATGCTGGGCCTGGATCGTCTCACC-3�, digestedwith NdeI, and inserted into the NdeI site of pVCHYN-4 (53). To constructCJW2959, pVsigpepCHYN-4 was delivered into CB15N by electroporation, andintegrants by single crossover into Pvan were selected. To construct CJW3049,pXGFPN-5 (53) was delivered into CJW2959 by conjugation, and integrants bysingle crossover into Pxyl were selected.

C. crescentus strains were grown in peptone-yeast extract (PYE) medium or indefined minimal medium (M2G) with shaking at 30°C (18), with appropriateantibiotics for selection when required. Agrobacterium tumefaciens and Sinorhi-zobium meliloti were grown in Luria Broth (LB) medium at 30°C. E. coli wasgrown in LB medium at 37°C. Adjustments of the pH of growth media were doneby adding Tris-HCl buffer (pH 8.0) or sodium phosphate buffer (pH 6.1) to afinal concentration of 20 mM (to create PYE-TRIS and LB-TRIS or PYE-P andLB-P, respectively). Xylose (0.3%) or vanillic acid (0.5 mM) was added to inducegene expression from the respective inducible promoters while glucose (0.2%)was used to repress the xylose-inducible promoter. Protein synthesis in strainCJW3049 was induced by addition of vanillic acid and xylose for 6 h and 4 h,respectively, prior to imaging. MreB inhibitors were added to the desired con-centrations from 1,000� stock solutions in dimethyl sulfoxide (DMSO).

Growth curve experiments were performed using a Bio-Tek Synergy 2 plate

TABLE 1. Strains and plasmids

Strain or plasmid Relevant genotype and/or description Source or reference

StrainsC. crescentus

CB15N Also known as NA1000; synchronizable variant of the wild-type strain CB15 19CJW2959 CB15N Pvan::pVsigpepCHYN-4 This studyCJW3049 CB15N Pvan::pVsigpepCHYN-4 Pxyl::pXGFPN-5 This studyJG5008 CB15N recA::Tn5 �mreB/pRK290/20R Pxyl-mreB; MreB depletion strain 20JS1003 CB15N rsaA::Kmr; does not produce an S-layer 65LS3814 CB15N Pxyl::pXGFP4-C1mreB; gfp-mreB under the control of Pxyl 22MT196 CB15N Pvan::pMT383; ftsZ-yfp under the control of Pvan 54YB1585 CB15N ftsZ::pBJM1; FtsZ depletion strain 66

E. coliDH5� Cloning strain PromegaFB9 PB103 �mreB/pFB112 (tet sdiA); stable mreB deletion strain 4MC1000 Wild-type E. coli strain 10PB103 Wild-type E. coli strain 14S17-1 RP4-2, Tc::Mu Km::Tn7; for plasmid mobilization 49

A. tumefaciens3101 Wild-type A. tumefaciens strain S. Dinesh-Kumar

S. meliloti1021 MB501; wild-type S. meliloti strain Bill Margolin

PlasmidspVCHYN-4 Integrating plasmid; contains Pvan-mCherry sequence; for vanillate-inducible

expression of mCherry53

pVsigpepCHYN-4 Integrating plasmid; contains Pvan-CC1996sigpep-mCherry; for vanillate-inducible expression of periplasm-exported mCherry

This study

pXGFPN-5 Integrating plasmid; contains Pxyl-gfp sequence; for xylose-inducibleexpression of GFP

53

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reader. In a 96-well plate, 5 �l of three or four independent overnight cultureswas inoculated into 195 �l of premade medium containing the desired amount ofdrugs or their solvent (DMSO). The plates were covered with a lid and incubatedat the appropriate temperature with shaking. Absorbance was read at 660 nm (C.crescentus) or 600 nm (A. tumefaciens, S. meliloti, and E. coli) every 2 min (C.crescentus, A. tumefaciens, and E. coli) or every 5 min (S. meliloti). For doublingtime and growth rate calculations, the reads from each well were individuallyplotted in a semilogarithmic scale, and the linear part of the graph was fitted toan exponential curve: y � aebt, where t is time. The doubling time (DT) wascalculated as DT � ln2/b and the growth rate r was calculated as r � 1/DT.

Drug synthesis. A22 and each analog were synthesized according to the fol-lowing general procedure. Substituted benzyl halide (1.5 Eq) was added to a 0.2M solution of thiourea in absolute ethanol. The reaction mixture was kept underrefluxing conditions for 16 h and then cooled down slowly to room temperature(RT). White crystals were isolated by filtration and washed twice with a cold 1:1diethyl ether-ethanol mixture. After extensive drying under reduced pressure,the carbamimidothioates were obtained in good to excellent yields (80 to 95%).The compound structures were verified by 1H and 13C nuclear magnetic reso-nance (NMR) (data not shown). For the synthesis of MP265, a solution of4-chlorobenzyl chloride (12.92 mmol; 2.08 g) and thiourea (13 mmol; 988 mg)was heated for 12 h in refluxing absolute ethanol (25 ml). After the solution wascooled down slowly to RT, 150 ml of diethyl ether was added slowly, and themixture was further cooled down to 0°C to induce crystallization of the product.After filtration, two washes with 40 ml of ice-cold diethyl ether, and extensivedrying under reduced pressure, 2-(4-chlorobenzyl)isothiouronium chloride wasisolated as white crystals (2.81 g; 94% yield). The structure of the product wasconfirmed using 1H and 13C NMR, and mass spectrometry (MS): 1H NMR (300MHz, D2O� ε d4-MeOD): � � 7.34 (s, 4H), 4.31(s, 2H); 13C NMR (75 MHz,D2O � ε d4-MeOD): � � 171.4, 134.8, 134.0, 131.5, 130.1, 35.5; MS (gas chro-matography [GC]/MS, electron impact [EI]): 158 (4-ClC6H4CHS�, 40%), 125(4-ClC6H4CH2

�, 100%). MP265 is also available commercially from variousvendors (Chemical Abstract Services [CAS] 544-47-8).

Light microscopy. Light microscopy imaging was performed at RT using 100�differential-interference contrast (DIC) or phase-contrast objectives on an NikonE1000 microscope equipped with a Hamamatsu Orca-ER LCD camera or aNikon Eclipse 80i microscope equipped with an Andor iXonEM� (DU-897E)electron-multiplying charge-coupled-device (EMCCD) camera and HamamatsuOrcaIIER camera. Cells were placed on 1 to 1.5% agarose pads made with M2G(for single-image acquisitions), with M2G containing 1% PYE (for fluorescencetime-lapse microscopy experiments), or with PYE (for DIC time-lapse experi-ments). For cell width measurements, the cells were fixed in Karnovski’s fixative(5% glutaraldehyde, 4% formaldehyde in 0.08 M sodium phosphate buffer, pH7.2) at RT for 1 h, stored at 4°C, and washed in phosphate-buffered saline (PBS)before imaging. This fixation procedure did not alter the shape or width of thecells (data not shown). Images were acquired and processed using Metamorphsoftware (Molecular Devices). Cellular widths were measured from phase-con-trast micrographs using a Matlab-based algorithm.

EM. For scanning electron microscopy (EM), 1 ml of culture was harvestedand centrifuged. Cells were resuspended in 500 �l of Karnovski’s fixative andstored at 4°C. Once all samples were collected, the fixed cells were washed twicein PBS and mounted onto poly-L-lysine-coated coverslips. The cells were dehy-drated by passage through increasing concentrations of ethanol, ending in 100%ethanol. Finally, the samples were critical point dried and gold coated beforebeing examined using a FEI XL-30 ESEM FEG microscope with the followingsettings: acceleration voltage, 10.0 kV; spot size, 3; working distance, 7.5 mm(17).

For transmission EM, the cells were pelleted, washed in PBS, and fixed for 1to 2 h in Karnovski’s fixative at RT. The cells were washed twice in PBS and fixedin aqueous 2% osmium tetroxide for 1 to 2 h at RT. Next, the cells were washedthree times in PBS, mixed with 2% agarose solution, and allowed to solidify at4°C. The agarose-trapped cells were dehydrated through an ascending series ofethanol solutions and embedded in EMbed812 (Electron Microscopy Sciences)according to the manufacturer’s instructions. In a variation of the procedure, theagarose-trapped cells were stained en bloc prior to the ethanol dehydration stepsby treating them for 2 h with a 1% uranyl acetate solution in 50% aqueousethanol (26). Ultrathin sections (40 to 100 nm) were cut with a diamond knife onan MT-1 Sorvall ultramicrotome, mounted onto copper grids, stained for 1 mineach with aqueous uranyl acetate and lead citrate, and visualized with a JEOLJEM-1230 transmission electron microscope operating at 80 kV. Digital imageswere acquired with a Hamamatsu ORCA-HR camera.

Isolation and analysis of peptidoglycan sacculi. PG sacculi were isolated aspreviously described (24, 39). Five hundred milliliters of cell culture in exponen-tial growth phase (optical density at 600 nm [OD660] of 0.2 to 0.5) was cooled for

10 min on ice and centrifuged at 10,400 � g for 10 min at 4°C. The pellet wasresuspended in 8 ml of ice-cold PBS and added dropwise under vigorous stirringto 8 ml of boiling 8% SDS. The mixture was boiled for 30 min with stirring whilewater was added throughout to maintain a constant volume. Sacculi were pel-leted at 440,000 � g in a Beckman Coulter Optima TLX Ultracentrifuge for 15min. The pellet was washed by sequential resuspension in water, followed byultracentrifugation until the supernatant was free of SDS as evaluated by theHayashi test (27). The sacculi were resuspended in 10 mM Tris-HCl buffer (pH7.0), incubated with 100 �g/ml amylase for 2 h at 37°C, followed by incubation for1 h with 200 �g/ml pronase at 60°C (pronase was preincubated for 2 h at 60°C),and boiled for 30 min in 4% SDS. The sacculi were washed free of SDS andstored at 4°C with 0.02% sodium azide in water. For transmission EM, the sacculisuspensions were diluted 1:5 in water, mounted onto glow-discharged Formvarcarbon-coated copper grids, stained with 1% uranyl acetate for 1 min, andvisualized similarly to the thin sections.

Muropeptide preparation and analysis. Muropeptide preparation wasachieved as described previously (24), with some modifications. Briefly, PGsacculi were isolated as described above and digested at 37°C overnight with 20�g/ml cellosyl in 20 mM sodium phosphate, pH 4.8. Cellosyl was removed byboiling the mixture at 100°C for 10 min and centrifugation at 15,000 � g for 30min. The muropeptide mixture was reduced with sodium borohydride in 0.25 M(final concentration) sodium borate, pH 9.0, for 30 min; then the pH wasadjusted to 3 to 4.5 using 20% phosphoric acid, and the samples were concen-trated under vacuum at RT in a Rotovap. The muropeptides were separated withan Agilent 1200 series HPLC system on a 250-mm by 4.6-mm, 3-�m C18 HypersylODS column. The solvent gradient was linear from 50 mM sodium phosphate,pH 4.31, to 70% (75 mM sodium phosphate, pH 4.95)–30% methanol in 135 min.The column was maintained at 55°C, and UV detection was performed at 205nm. Fractions corresponding to peaks of interest were collected manually andanalyzed by mass spectrometry as previously described (6). All muropeptide-related calculations were performed as previously described (24).

RESULTS

A22 can have an MreB-independent toxic effect on bacterialgrowth. A22 has been used to reversibly disrupt the localiza-tion and function of MreB in C. crescentus and other Gram-negative bacteria (23, 32, 41, 45, 52). However, A22 is knownto reduce the growth rate of an E. coli mutant that lacks MreBat concentrations of 50 �g/ml (212 �M) and higher (35), sug-gesting that A22 inhibits cellular function(s) other than that ofMreB. Given the rapidly increasing use of A22 in bacterialstudies, we tested whether this potentially toxic effect of A22varied with the bacterial species and/or growth conditionsused. We first examined the effects of different concentrationsof A22 on the growth of three rod-shaped alphaproteobacteria:C. crescentus, S. meliloti, and A. tumefaciens. S. meliloti and A.tumefaciens do not have an MreB homolog (11) while MreB ispresent and essential for viability in the closely related C.crescentus (20, 22). We found that A22 affected the rate andextent of growth of all three species in a concentration-depen-dent manner (Fig. 1A and 2; see also Table S1 in the supple-mental material). A culture medium effect was also observed asthe growth-inhibitory effect was significantly more pronouncedwhen C. crescentus was grown in minimal M2G medium thanwhen it was grown in rich PYE medium (Fig. 1A and 2).Treatment with the commonly used concentration of 10 �g/ml(50 �M) of A22 reduced the growth rates not only of C.crescentus but also of A. tumefaciens and S. meliloti (Fig. 1Aand 2). Consistent with the absence of mreB from the genomesof these last two species, their rod morphologies were notaffected by A22 treatment (data not shown). A �mreB E. colistrain (FB9) and its wild-type parent strain (PB103) also ex-hibited slower growth rates in an A22 concentration-depen-dent fashion (Fig. 1A and 2 and Table S1), confirming the

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FIG. 1. Inhibitory effects of A22 and MP265 on bacterial growth. C. crescentus CB15N, A. tumefaciens 3101, S. meliloti 1021, and E. coliPB103 and FB9 (PB103 �mreB) were grown in the presence of the noted concentrations of A22 or MP265, and the optical densities of thecultures were recorded. Cells were grown in the media indicated on the figure. Each curve represents the average optical density of at leastthree independent cultures.

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previous study (35). E. coli was, however, considerably moreresistant than C. crescentus, A. tumefaciens, or S. meliloti. Theseresults argue that A22 has an MreB-independent toxic effecton bacterial growth that varies among species. This toxicity isconsistent with the finding that A22 binds to MreB as a low-affinity ADP mimic (dissociation constant [Kd] of 1.32 �M)(3) and may therefore affect other nucleotide-binding pro-cesses.

MP265 (4-chlorobenzyl carbamimidothioate) is an A22 an-alog with reduced toxicity on cell growth. As A22 affectedMreB-independent processes, we searched for other drugs thatwere more specific in disrupting MreB function. Sceptrin, a nat-urally occurring compound that was shown to pull down MreB invitro (46), did not disrupt the rod shape of C. crescentus when usedat 1 or 5 �g/ml (1.6 or 8.1 �M, respectively) and completelyblocked cell growth at 5 �g/ml (data not shown).

Structural analogs of A22 have been shown to induce cellrounding in E. coli by presumably acting on MreB (30, 31). Wetherefore synthesized and tested a small library of A22 analogsin assays designed to evaluate the anti-MreB effects of thesecompounds (see Table S2 in the supplemental material). Wescored their ability to disrupt green fluorescent protein (GFP)-MreB localization and to induce cell rounding when the drugswere added at a concentration of 50 �M to a growing cultureof wild-type C. crescentus cells (Table S2). Under these exper-imental conditions, A22-treated cells displayed a largely diffuseGFP-MreB localization and adopted a lemon-shaped mor-phology (23). Of the A22 analogs tested, MP265 and MP253exhibited anti-MreB activities comparable with those of A22

(Table S2). As MP265 appeared to be less toxic than eitherA22 or MP253 in preliminary growth curve experiments (datanot shown), we decided to further characterize its effects onMreB and bacterial growth.

In C. crescentus, MreB localization alternates betweenpatchy/helical and midcell ring patterns during the cell cycle(20, 22). Asynchronous cell populations treated with only thedrug solvent (DMSO) for 1 h displayed the expected patchy/banded localization of GFP-MreB (Fig. 3A, Untreated), aspreviously reported (22). Under similar conditions, 5 �M A22or MP265 caused partial delocalization of the GFP-MreB sig-nal (Fig. 3A), and at 50 �M (Fig. 3A), both drugs delocalizedmost of the GFP-MreB signal although some weak GFP-MreBbands could still be observed in some cells, as previously notedfor A22 (1). We concluded that MP265 is equivalent to A22 inits ability to disrupt MreB filaments in live C. crescentus cells.However, MP265 has the advantage of being less toxic thanA22. Invariably, higher concentrations of MP265 than of A22were needed to reduce growth rates and to increase the gen-eration times of the tested bacterial cultures (Fig. 1B and 2; seealso Table S1 in the supplemental material). When used at 50�M, a commonly employed concentration for A22 treatment,MP265 barely decreased the growth rates of the MreB-lackingbacteria A. tumefaciens and S. meliloti, and even at 500 �M,MP265 had no detectable inhibitory effect on the growth of thetested �mreB E. coli strain under our experimental conditions(Fig. 2 and Table S1).

Basic pH of the growth medium enhances the rod shape-disrupting effect of A22 and MP265. Treatment with A22 or

FIG. 2. Effects of A22 and MP265 on bacterial growth rates. The growth rate of each bacterial culture grown in the presence of a 0, 5, 25, 50,100, 250, or 500 �M concentration of either A22 or MP265 was calculated from the exponential part of each individual growth curve (shown inFig. 1), and divided by the average growth rate of the respective culture in the absence of any drug. The obtained relative growth rates were plottedas a function of the drug concentration added to the culture (average standard deviation of at least three independent values).

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MP265 in PYE medium resulted in a gradual widening of C.crescentus cells over time (Fig. 3B; see also Fig. S1 in thesupplemental material). At 5 �M, both drugs were particularlypotent in disrupting rod cell morphology in denser cultures,and this effect was correlated with an increase in the pH of theculture medium during growth (data not shown). Sterile PYEmedium had a pH of 6.6 to 6.8 while medium from stationary-phase cultures of C. crescentus had a pH of 8.5. To eliminatethe effect of pH variation during growth, we buffered the me-dium at pH 7.8 (PYE-TRIS medium). Treatment of PYE-TRIS cultures with either A22 or MP265 at 5 �M resulted inthe most significant increases in cell widths (Fig. 3B and S1),confirming an enhancement of drug activity by basic pH similar

to that documented for other amine-containing drugs acting oncytoplasmic targets (28). This effect was likely due to increasedconcentration of the membrane-diffusible, nonprotonatedform of the drug in the basic medium, leading to increasedintracellular accumulation of the drug. Conversely, treatmentwith a 5 �M concentration of either drug in medium bufferedat an acidic pH of 6.6 (PYE-P) caused no significant increasein cell widths (data not shown). The pH dependence of the rodshape-disrupting effect of 5 �M A22 also occurred with E. colicells (see Fig. S2 in the supplemental material). Treatment ofPYE-TRIS C. crescentus cultures with 50 �M MP265 or A22produced smaller cell width increases than treatment with 5�M, with A22 being less potent (Fig. 3B and S1). This effect

FIG. 3. Disruptive effects of A22 and MP265 treatments on GFP-MreB localization and rod morphology of C. crescentus. (A) Synthesis ofGFP-MreB was induced for 2 h in cultures of strain LS3814; then the cells were treated for 1 h with the indicated drug concentrations whileinduction was maintained, placed on agarose pads containing the same drug concentrations used for treatment, and imaged by light microscopy.Treatment with the 5 �M drug concentrations was performed in PYE-TRIS medium while treatment with the 50 �M drug concentrations wasperformed in PYE medium. (B) Wild-type CB15N cultures growing in PYE or PYE-TRIS were treated with the indicated drug concentrations.Cells harvested before the addition of the drugs and after 3 and 6 h of growth in the presence of the indicated amounts of drugs were fixed andimaged by phase-contrast microscopy. The maximum width of each cell was measured using a Matlab-based algorithm, and the mean width of 400to 2,000 cells from each culture was determined. The plot depicts the average of the mean cell width from three independent experiments for eachtreatment condition as a function of the duration of the treatment.

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was likely due to enhanced growth inhibition by A22 andMP265 under these treatment conditions, with A22 being moretoxic than MP265 (Fig. 3B, S1, and S3 and Table S3). Cellwidening induced by MreB inhibition is growth dependent(23). At 5 �M, A22 or MP265 did not affect the growth ofbacteria lacking MreB when bacteria were grown in the basicmedium LB-TRIS (see Fig S3 and Table S3 in the supplemen-tal material), prompting us to use a 5 �M concentration ofeither MP265 or A22 on C. crescentus growing in PYE-TRIS(pH 7.8) to disrupt rod shape in all of the following experi-ments.

Morphological changes associated with recovery of C. cres-centus cells from A22 or MP265 treatment comprise cell thin-ning, filamentation, branching and shedding of outer mem-brane vesicles. After 4 h of treatment with 5 �M MP265 inPYE-TRIS medium, the cells acquired wide, lemon-like shapes(Fig. 4Aii), occasionally presenting a bump on one lateral side.To monitor their recovery after dug removal, lemon-shaped,MP265-treated cells were washed and imaged by time-lapseDIC microscopy. Immediately after removal of the drug, afraction of the lemon-shaped cells gradually became thinneruntil they reached normal cell widths (see Movie S1 in thesupplemental material). The cell-thinning process was strikingas it occurred along the entire lateral wall of lemon-shaped C.crescentus cells, including where the cells were originally wide.Cell thinning was also accompanied by cell elongation, produc-ing long filamentous cells (Movie S1). Other cells within thepopulation also began to thin and elongate but eventually lysedon the agarose growth pad (Movie S2). Thinning and elonga-tion occurred concomitantly (see Fig. S4 in the supplementalmaterial). These dramatic morphology changes required denovo PG synthesis since treatment with the PG synthesis in-hibitor fosfomycin prevented cell shape recovery after removalof the MreB inhibitor (data not shown).

The processes of cell thinning and filamentation were alsoaccompanied by abundant shedding of cellular particles fromthe sides of the growing cells (Movies S1 and S2, arrowheads).We obtained similar results with cells recovering from A22treatment (data not shown). Cell thinning, filamentation, andshedding of cellular particles could also be observed when theMP265-treated cells were washed and allowed to grow in theabsence of the drug in liquid culture, as shown by scanning EM(Fig. 4A). Untreated and MP265-treated C. crescentus cellshad curved-rod and lemon-like shapes, respectively, andlargely smooth cell surfaces (Fig. 4Ai and ii; see also Fig. S5iand ii in the supplemental material). After removal of the drug,the lemon-shaped cells became increasingly thinner and longer(Fig. 4Aiii to vii and S5iii to vii). Their surfaces became pro-gressively more irregular, and extensive blebbing could be ob-served on the surfaces of many of the recovering cell filaments(Fig. 4Aiv and v and S5iv to vii). By the time the cells hadreached widths similar to those of wild-type cells, fractions ofthe cell population had grown cell branches or ectopic stalkspositioned along the side wall (18% and 10%, respectively; n �108) (Fig. 4Avi to vii and S5vi). These phenotypes are remi-niscent of cells recovering from MreB depletion, as previouslydescribed (64).

A surprising and previously unreported phenotype associ-ated with the recovery process was the profuse shedding ofcellular vesicles. We also observed this shedding in cells recov-

ering from MreB depletion (see Movie S3 in the supplementalmaterial), indicating that it is not due to destabilization of theouter membrane by MP265 or A22 independent of their actionon MreB. To identify the contents of the shedding particles, weconstructed a C. crescentus strain (CJW3049) (see Materialsand Methods) that produces the fluorescent proteins GFP andmCherry in the cytoplasm and periplasm, respectively. Lemon-shaped, MP265-treated CJW3049 cells growing after drug re-moval elongated, became thinner, occasionally grew smallbranches, and shed cellular particles (Fig. 4B and Movie S4), asexpected. The cytoplasmic GFP signal remained containedwithin the cell body (including the branches) throughout therecovery process and was completely absent from the shedparticles, suggesting that they do not contain cytoplasm (Fig.4B and Movie S4). The periplasmic mCherry signal, on theother hand, became concentrated at distinct locations whereshedding occurred and formed distinct bright foci that colocal-ized with the shed particles (Fig. 4B and Movie S4). That themCherry signal was higher in the shed vesicles than in thenonblebbing cell periphery is likely due to the large volume ofperiplasm enclosed within vesicles relative to the rest of theperiplasm. These observations strongly suggest that the shedparticles are large outer membrane vesicles (OMVs).

We confirmed this result by using transmission EM of ultra-thin sections through fixed, plastic-embedded cells. Under ourfixation and imaging conditions, the outer membrane (OM)appeared frizzled, and the S-layer was not visible (see Fig. S6in the supplemental material), consistent with previous reports(42, 51). The peptidoglycan layer was particularly well stainedin our preparations, and the inner membrane was easier todetect (Fig. S6) when we used a uranyl acetate en bloc stainingstep prior to the ethanol dehydration procedure (26). Trans-mission EM images of ultrathin sections of cells recoveringfrom treatment with A22 or MP265 revealed that the sheddingparticles were bound by a single-layered structure continuouswith the outer membrane of the cell, confirming that they areOMVs (Fig. 4C and data not shown).

Given the filamentation defect associated with recoveryfrom drug treatment, we examined the localization of the ma-jor cell division protein FtsZ. Fluorescence microscopy of astrain producing FtsZ fused to yellow fluorescent protein(FtsZ-YFP) showed that FtsZ formed a ring (which appears asa band) in untreated cells but failed to form a ring in lemon-shaped A22-treated cells and, instead, largely localized in atight focus either within the bumps observed by DIC micros-copy or at one pole of the cells (see Fig. S7A in the supple-mental material). The bumps were also noticeable in PG sac-culi isolated from A22-treated cells (Fig. S7B, arrowhead). AsFtsZ-driven cell division results in the synthesis of new poles indaughter cells, it is conceivable that the observed local con-centration of FtsZ on the side of lemon-shaped A22-treatedcells drives local PG synthesis, resulting in the creation of anectopic pole. This was supported by the observation thatbumps occasionally developed into cell branches (see MovieS5, arrow, in the supplemental material). Shortly after removalof A22, FtsZ-YFP formed a ring in the recovering cells; how-ever, the ring showed instability and moved along the elongat-ing filament until late into the recovery process when stableFtsZ rings drove division of the filament (Movie S5).

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FIG. 4. Phenotypes associated with recovery from MP265 and A22 treatment. (A) Scanning EM images of CB15N cells before MP265treatment (i); after 4 h of treatment with 5 �M MP265 in PYE-TRIS (ii); or at 2 h (iii), 4 h (iv), and 6 h (v to vii) after removal of MP265.(B) Selected images of CJW3049 cells from a time-lapse sequence of recovery from 4 h of 5 �M MP265 treatment. Synthesis of cytoplasmic GFPand periplasm-exported mCherry was achieved by growing cells in the presence of 0.3% xylose and 0.5 mM vanillic acid for 4 h and 6 h, respectively,prior to imaging on agarose pads containing xylose and vanillic acid. Time is given as hours:minutes. (C) Transmission EM images of CB15N cellsafter 6 h of recovery from 4 h of 5 �M A22 treatment. OM, outer membrane.

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The morphology of the peptidoglycan cell walls changesduring recovery from A22 treatment. Bacterial cell morphol-ogy is thought to be determined by the shape of the pep-tidoglycan cell wall. As recovery from A22 or MP265 treatmentwas accompanied by extensive cell shape remodeling, we in-quired whether any changes in the peptidoglycan accompaniedthis process. Invariably, isolated PG sacculi retained the shapeof the cells from which they had been isolated (Fig. 5; note thatthe spheres in the EM images represent polyhydroxybutyrate[PHB] storage granules trapped inside the PG mesh) (43).Uranyl acetate staining of sacculi isolated from untreated orA22-treated cells resulted in a uniform, light stain (Fig. 5A andB). Sacculi isolated from cells early during the process ofrecovery from A22 treatment (at 1, 2, and 3 h after removal ofA22) and stained with uranyl acetate under identical condi-tions invariably presented darker-stained striations thatcrossed the otherwise lightly stained sacculi along their shortaxes (Fig. 5C to E). These striations were most dense in images

of sacculi isolated 1 h after removal of A22 (Fig. 5C). As thecells progressed through the recovery process and the sacculibecame thinner, the striations were spread farther apart andwere fainter (Fig. 5C to E). They were completely absent fromsacculi that had reached normal widths (Fig. 5F) (5 h afterremoval of A22). These observations suggested that the stria-tions might represent changes in the PG cell walls associatedwith, and presumably important for, the recovery of the rodshape. Importantly, these dark striations were distinct fromregular folds of the sacculi that can occur as the sacculi laydown on the EM grids (see Fig. S8 in the supplemental mate-rial) (16).

As noted above, not all cells in a population were able tofully recover after removal of A22 or MP265. Consistent withthis, a fraction of the sacculi from the samples harvested at 3 or5 h into the recovery process remained aberrantly wide andretained an elongated lemon shape (11% of sacculi [n � 341]for 3 h and 5.5% of sacculi [n � 475] for 5 h). These sacculi

FIG. 5. Transmission electron micrographs of C. crescentus sacculi isolated at different time points during recovery from A22. (A) Untreatedcells. (B) Cells treated for 4 h with 5 �M A22 in PYE-TRIS. (C to E) Cells after 1 h (C), 2 h (D), and 3 h (E) of recovery from a 4-h treatmentwith 5 �M A22 in PYE-TRIS. (F) Cells after 5 h of recovery from a 5-h treatment with 5 �M A22 in PYE-TRIS. (G) High-magnification imageof a sacculus containing strongly stained striations, isolated after 3 h of recovery from a 4-h treatment with 5 �M A22 in PYE-TRIS. PHB,polyhydroxybutyrate granule trapped inside the sacculus.

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invariantly displayed thick, dark-stained, discrete striationsalong the length of the sacculus (Fig. 5G; see also Fig. S9 in thesupplemental material). These striations were generallyslightly tilted with respect to the short axis of the sacculi andoften connected with other striations, forming V-like struc-tures that crossed each other along the length of the sacculi(Fig. 5G and S9).

Changes in PG composition during recovery from A22 treat-ment. Given that A22 treatment and recovery from A22 treat-ment involve extensive changes in the morphology of PG sac-culi, we determined whether these changes were associatedwith changes in PG composition. The muropeptide composi-tion of the C. crescentus PG has only been roughly describedpreviously (39). We therefore used an HPLC-based protocol toseparate and quantify the reduced muropeptide buildingblocks of the C. crescentus PG (24, 39). In order to unambig-uously assign molecular structures to the separated muropep-tides, we collected the fractions containing the major mu-ropeptide peaks and analyzed them by MS (6). The overallmuropeptide profile of C. crescentus differed significantly fromthat of E. coli (24, 25) although the PG type was the same andtypical for Gram-negative bacteria, and major muropeptideswith the same structure were present in both species (Fig. 6A;see also Table S4 in the supplemental material). In contrast toE. coli PG, C. crescentus PG did not contain detectableamounts of covalently linked Braun’s lipoprotein (Lpp) (39,43). This is consistent with the absence from the C. crescentusgenome of the gene encoding this outer membrane anchor.

Second, a variant of the normal pentapeptide with Gly replac-ing the terminal D-Ala [penta(Gly5)] was highly abundant inboth free and cross-linked forms, consistent with previouslypublished data (39) (Table 2 and Fig. 6B). Third, cross-linkedmuropeptides were more abundant, including trimers and tet-ramers, leading to a high overall degree of cross-linkage of theC. crescentus PG (34%) (Table 2 and Fig. 6B) compared to thatof E. coli PG (23%) (24) (note that the degree of cross-linkageis a numerical value and is not the same as the molar percent-age of peptides present in cross-links). We also identified byMS a muropeptide dimer carrying an L,D-cross-link (m-Dap-mDap) (Fig. 6A and Table S4). L,D-Cross-links were thought tobe absent from the C. crescentus PG (39); their presence, how-ever, is not surprising, given that the C. crescentus genome en-codes a putative L,D–transpeptidase (locus cc_1511; BLASTPvalue of 4.4 � e�33 against the E. coli K12 L,D-transpeptidaseYcbB) (38). Fourth, 1,6-anhydro-MurNAc glycan chain endswere abundant (Table 2 and Table S5), including as part of di-anhydro cross-linked species, which are extremely rare in E. coli(24, 25). This correlated with average glycan chain lengths (9disaccharide subunits/glycan chain) (Table 2) that were signifi-cantly shorter than those of E. coli (30 disaccharide subunits/glycan chain) (24).

Having identified the major muropeptides present in the C.crescentus PG, we then characterized and compared the mu-ropeptide composition of PG isolated from untreated cells,A22-treated cells, and cells recovering from A22 treatment(see Table S5 in the supplemental material). A22 treatment

FIG. 6. Muropeptide composition of C. crescentus PG during A22 treatment and recovery from A22 treatment. (A) UV chromatogram ofHPLC-separated muropeptides isolated from the PG of CB15N cells. Peak identities determined by MS/MS: Tri, GlcNAc-MurNAc-L-Ala-L-�-Glu-m-Dap; tetra, Tri-D-Ala; Penta, Tetra-D-Ala; Penta(Gly5), Tetra-Gly; Anh, 1,6-anhydro-MurNAc; (D,L), m-Dap-m-Dap cross-link. (B) Com-position (in molar percentage) of various muropeptide species in the PG of untreated CB15N cells (mean values standard deviation ofmeasurements from three independent samples). (C) Changes in the composition of C. crescentus PG during A22 treatment and recovery fromtreatment. The molar percentage of each species in a given sample is expressed as a percentage of the average molar percentage of the speciesin the PG from untreated cells. Treatment conditions are as indicated on the figure. Bars represent mean values standard deviation of three (U),two (T, 1, and 2), and one (3) independent PG samples.

TABLE 2. Summary of C. crescentus PG composition during A22 treatment and recovery from treatment

Muropeptide species or feature

PG composition (molar % SD) of cells

Withouttreatmenta

With A22treatmentb

During recovery at:

1 hb 2 hb 3 hc

Monomer 41.54 1.23 40.39 0.40 36.20 0.15 38.81 0.03 38.57Dimer 38.68 1.35 38.04 0.89 47.77 0.14 46.96 0.03 46.60Trimer 17.23 0.69 17.98 1.31 14.48 0.56 12.70 0.18 13.10Tetramer 4.10 0.40 6.10 0.15 2.91 0.31 2.14 0.15 2.23Cross-linkaged 33.89 0.72 35.59 0.55 35.72 0.21 33.55 0.02 33.71Pentapeptide (Ala5) 20.33 0.44 19.01 0.37 18.31 0.07 17.91 0.05 19.18Pentapeptide (Gly5) 14.70 1.08 14.94 1.18 22.21 0.04 23.13 0.18 21.73Pentapeptide (Total) 35.03 1.33 33.94 0.81 40.51 0.12 41.04 0.23 40.91Tetrapeptide 63.75 1.05 64.86 1.06 58.53 0.17 58.03 0.48 58.18Tripeptide 1.09 0.56 1.06 0.26 0.90 0.04 0.89 0.25 0.85Anhydro 10.66 0.50 13.50 0.005 6.46 0.34 5.28 0.15 5.66Average glycan chain lengthe 9.40 0.45 7.41 0.003 15.50 0.83 18.96 0.55 17.66

a Results of three independent experiments.b Results of two independent experiments.c Results of one experiment.d Calculated as in described in reference 26.e Values represent number of disaccharide subunits/glycan chain.

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caused a 20% decrease in the average glycan chain length(Table 2 and Fig. 6C). A similar decrease of the average glycanchain length has been observed in the PG of A22-treated E.coli cells (57). The amount of highly cross-linked tetramericmuropeptides was increased by about 50% after A22 treatment(Table 2 and Fig. 6C). The degree of cross-linkage, however,was not significantly changed in A22-treated cells compared tountreated cells (Table 2 and Fig. 6C). This suggests that theA22-induced loss of the rod morphology was not due to reduc-tion of the number of PG cross-links.

Recovery from A22 treatment was accompanied by consid-erable changes in PG composition, pointing toward enhancedPG synthesis. Most importantly, the average glycan chainlength was increased up to 2.5-fold compared to the glycanchain length of A22-treated cells and up to 2-fold compared tothe value obtained from untreated cells (Table 2 and Fig. 6C).The amount of un-cross-linked, glycine-containing pentapep-tide, penta(Gly5), in the muropeptide side chains was in-creased by 50% compared to the amounts of the same spe-cies in PG from untreated or A22-treated cells. Similarly,recovery from A22 treatment was accompanied by 20% in-crease in the amount of dimeric muropeptides, which are di-rectly formed in the transpeptidation reactions that cross-linknewly inserted precursors to existent PG. Conversely, trimericand tetrameric muropeptides, which are formed in reactions ofpentapeptides with previously formed dimeric and trimericacceptors, were decreased by at least 20% and 50%, respec-tively, in PG of cells recovering from A22 treatment comparedto PG from A22-treated cells (Table 2 and Fig. 6C). Recoveryof the rod shape following A22 treatment is therefore accom-panied by extensive remodeling of the PG at both morpholog-ical and structural levels.

DISCUSSION

We show here that the widely used MreB inhibitor A22affects bacterial growth by also acting on an essential target(s)other than MreB. This MreB-independent toxic effect variesdepending on the species subjected to treatment and on thetreatment conditions used (e.g., medium and pH). Our resultsare consistent with previous documentation of MreB-indepen-dent toxic effects of A22 on the growth of E. coli (35) or of A22structural analogs on the growth of the Gram-positive Staph-ylococcus aureus (31), which lacks MreB (11). The species-dependent variation in toxicity levels requires careful exami-nation of the growth-inhibitory effect of A22 or its analogswhenever such drugs are used in the study of a new bacterialspecies. Furthermore, the discovery of MP265 as a compoundwith anti-MreB activity similar to that of A22, but with lowertoxicity, provides a better tool for reversible disruption ofMreB function in Gram-negative bacteria. Another importantconsideration is the pH of the culture medium since it has asignificant effect on the activity and toxicity of both A22 andMP265.

Cells recovering from A22 or MP265 treatment thinned untilthey reached normal widths. Such a remarkable change in cellwidth implies extensive PG remodeling within the wide sacculiof MreB-disrupted cells. Transient, dark, uranyl acetate-stained striations on sacculi appeared during this thinning pro-cess. In nonrecovering cells, the striations were exacerbated

and displayed clear V-shaped patterns, which may be related tothe suggested helical localization of MreB filaments in livecells. The projection of three-dimensional helices on a planarsurface, such as the EM grids, would resemble crossing Vs.These striations were present only in sacculi from cells recov-ering from A22 treatment which had not yet returned to nor-mal widths. They were also distinct from previously describedfolded PG (16), and they displayed highly regular patterns. It istherefore highly unlikely that they are folding or staining arti-facts. Previously, thicker, multilayered PG found in the septaof PG hydrolase mutants of E. coli was shown to also staindarker with uranyl acetate than the rest of the sacculi (44).Thus, while we cannot completely rule out other explanations,we believe that the striations represent areas of thicker, mul-tilayered PG, likely where new insertion took place. We as-sume that the disappearance of the striations from sacculi atlate time points of recovery is due to the action of PG hydro-lases. An endopeptidase activity, for example, could accountfor the observed disappearance of the striations and concom-itant decrease in the amounts of trimeric and tetrameric mu-ropeptides and increase of the dimeric species (Fig. 6C). Thetransient presence of V-shaped striations during rod shaperecovery correlates with a considerable increase in glycan chainlength. PG synthesis is thought to occur along helical pathswithin the side wall (15, 59), and the glycan strands are thoughtto be oriented similarly (21, 61). Our results are consistent withthis model, and the longer glycans found in the PG of recov-ering cells may be present within the striations and may berequired to reduce the width of the wide sacculi. Conversely,loss of rod shape following MreB disruption may be due to lossof transverse/helical orientation of newly inserted glycanstrands.

Our findings that MreB inactivation results in a decrease ofthe glycan strand length and that restoration of MreB functionresults in a considerable increase of the glycan length suggestthat MreB may either promote the glycan strand-synthesizingactivity of the PBPs, inhibit the strand-cutting activity of thelytic transglycosylases, or both. A careful biochemical analysisof PG synthesis in E. coli has shown that MreB inactivation byA22 significantly inhibits the synthesis of new glycan strandsand causes accumulation of the soluble UDP-MurNAc-pen-tapeptide PG precursor in the cytoplasm (55). The same studyalso demonstrated that lytic transglycosylase activity was notaffected by A22 treatment, and another study showed that A22treatment causes a decrease of the average glycan strandlength in E. coli, consistent with our observations in C. cres-centus (55, 57). These corroborated findings argue that MreBpromotes, directly or indirectly, PBP transglycosylase activity.This function may be achieved via protein-protein interactionswithin the elongase complex or through regulation of the de-livery of PG precursors to periplasmic PBPs. MreB may alsoaffect PBP activity and PG growth by altering the mechanicalproperties of the cell wall (13).

The shedding of OMVs was another surprising phenotypeassociated with recovery from A22 or MP265 treatment. Adifference between the growth rate of the outer membrane andthat of the PG during recovery may be responsible for anaccumulation of excess outer membrane and blebbing. An-other, perhaps more likely, possibility is that the intense PGremodeling associated with recovery of rod morphology causes

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the loss of PG material that served as points of anchorage forouter membrane proteins. Local detachment of the outermembrane from the PG layer would favor blebbing. Indeed, inSalmonella, loss of attachments between PG and outer mem-brane results in increased shedding of OMVs (12), consistentwith a proposed model for the formation of such vesicles (40).

ACKNOWLEDGMENTS

We are grateful to L. Shapiro, P. de Boer, Y. Brun, J. Smit, J. Gober,S. Dinesh-Kumar, and W. Margolin for providing strains and plasmids;to B. Piekos, M. Mooseker, and Z. Jiang for support with the EM; toK. Bui and J. Gray for support with HPLC and tandem MS (MS/MS)analyses; to O. Sliusarenko for the Matlab-based software; and to H.Lam and the members of the Jacobs-Wagner laboratory for criticalreading of the manuscript.

This work was supported by the National Institutes of Health(GM076698 and GM065835 to C.J.-W.) and the European Commis-sion within the EUR-INTAFAR project (to W.V.). C.N.T. was sup-ported in part by a Yale College Dean’s Office Science, Technologyand Research Scholars (STARS) II fellowship; M. P. was supported bya Human Frontier Science Program Organization for a Cross Disci-plinary Fellowship, and S.P. was supported by a PEW Latin Ameri-can Fellowship. C.J.-W. is a Howard Hughes Medical Instituteinvestigator.

REFERENCES

1. Aaron, M., G. Charbon, H. Lam, H. Schwarz, W. Vollmer, and C. Jacobs-Wagner. 2007. The tubulin homologue FtsZ contributes to cell elongation byguiding cell wall precursor synthesis in Caulobacter crescentus. Mol. Micro-biol. 64:938–952.

2. Alyahya, S. A., R. Alexander, T. Costa, A. O. Henriques, T. Emonet, and C.Jacobs-Wagner. 2009. RodZ, a component of the bacterial core morpho-genic apparatus. Proc. Natl. Acad. Sci. U. S. A. 106:1239–1244.

3. Bean, G. J., S. T. Flickinger, W. M. Westler, M. E. McCully, D. Sept, D. B.Weibel, and K. J. Amann. 2009. A22 disrupts the bacterial actin cytoskeletonby directly binding and inducing a low-affinity state in MreB. Biochemistry48:4852–4857.

4. Bendezu, F. O., and P. A. de Boer. 2008. Conditional lethality, divisiondefects, membrane involution, and endocytosis in mre and mrd shape mu-tants of Escherichia coli. J. Bacteriol. 190:1792–1811.

5. Bendezu, F. O., C. A. Hale, T. G. Bernhardt, and P. A. de Boer. 2009. RodZ(YfgA) is required for proper assembly of the MreB actin cytoskeleton andcell shape in E. coli. EMBO J. 28:193–204.

6. Bui, N. K., J. Gray, H. Schwarz, P. Schumann, D. Blanot, and W. Vollmer.2009. The peptidoglycan sacculus of Myxococcus xanthus has unusual struc-tural features and is degraded during glycerol-induced myxospore develop-ment. J. Bacteriol. 191:494–505.

7. Cabeen, M. T., and C. Jacobs-Wagner. 2007. Skin and bones: the bacterialcytoskeleton, cell wall, and cell morphogenesis. J. Cell Biol. 179:381–387.

8. Carballido-Lopez, R., and J. Errington. 2003. The bacterial cytoskeleton: invivo dynamics of the actin-like protein Mbl of Bacillus subtilis. Dev. Cell4:19–28.

9. Carballido-Lopez, R., A. Formstone, Y. Li, S. D. Ehrlich, P. Noirot, and J.Errington. 2006. Actin homolog MreBH governs cell morphogenesis bylocalization of the cell wall hydrolase LytE. Dev. Cell 11:399–409.

10. Casadaban, M. J., and S. N. Cohen. 1980. Analysis of gene control signals byDNA fusion and cloning in Escherichia coli. J. Mol. Biol. 138:179–207.

11. Daniel, R. A., and J. Errington. 2003. Control of cell morphogenesis inbacteria: two distinct ways to make a rod-shaped cell. Cell 113:767–776.

12. Deatherage, B. L., J. C. Lara, T. Bergsbaken, S. L. Rassoulian Barrett, S.Lara, and B. T. Cookson. 2009. Biogenesis of bacterial membrane vesicles.Mol. Microbiol. 72:1395–1407.

13. de Boer, P. A. 2009. Bend into shape. EMBO J. 28:1193–1194.14. de Boer, P. A., R. E. Crossley, and L. I. Rothfield. 1988. Isolation and

properties of minB, a complex genetic locus involved in correct placement ofthe division site in Escherichia coli. J. Bacteriol. 170:2106–2112.

15. den Blaauwen, T., M. A. de Pedro, M. Nguyen-Disteche, and J. A. Ayala.2008. Morphogenesis of rod-shaped sacculi. FEMS Microbiol. Rev. 32:321–344.

16. de Pedro, M. A., J. C. Quintela, J. V. Holtje, and H. Schwarz. 1997. Mureinsegregation in Escherichia coli. J. Bacteriol. 179:2823–2834.

17. Ebersbach, G., A. Briegel, G. J. Jensen, and C. Jacobs-Wagner. 2008. Aself-associating protein critical for chromosome attachment, division, andpolar organization in Caulobacter. Cell 134:956–968.

18. Ely, B. 1991. Genetics of Caulobacter crescentus. Methods Enzymol. 204:372–384.

19. Evinger, M., and N. Agabian. 1977. Envelope-associated nucleoid from Cau-lobacter crescentus stalked and swarmer cells. J. Bacteriol. 132:294–301.

20. Figge, R. M., A. V. Divakaruni, and J. W. Gober. 2004. MreB, the cellshape-determining bacterial actin homologue, co-ordinates cell wall mor-phogenesis in Caulobacter crescentus. Mol. Microbiol. 51:1321–1332.

21. Gan, L., S. Chen, and G. J. Jensen. 2008. Molecular organization of Gram-negative peptidoglycan. Proc. Natl. Acad. Sci. U. S. A. 105:18953–18957.

22. Gitai, Z., N. Dye, and L. Shapiro. 2004. An actin-like gene can determine cellpolarity in bacteria. Proc. Natl. Acad. Sci. U. S. A. 101:8643–8648.

23. Gitai, Z., N. A. Dye, A. Reisenauer, M. Wachi, and L. Shapiro. 2005. MreBactin-mediated segregation of a specific region of a bacterial chromosome.Cell 120:329–341.

24. Glauner, B. 1988. Separation and quantification of muropeptides with high-performance liquid chromatography. Anal. Biochem. 172:451–464.

25. Glauner, B., J. V. Holtje, and U. Schwarz. 1988. The composition of themurein of Escherichia coli. J. Biol. Chem. 263:10088–10095.

26. Graham, L., and J. M. Orenstein. 2007. Processing tissue and cells fortransmission electron microscopy in diagnostic pathology and research. Nat.Protoc. 2:2439–2450.

27. Hayashi, K. 1975. A rapid determination of sodium dodecyl sulfate withmethylene blue. Anal. Biochem. 67:503–506.

28. Hille, B. 1977. The pH-dependent rate of action of local anesthetics on thenode of Ranvier. J. Gen. Physiol. 69:475–496.

29. Holtje, J. V. 1998. Growth of the stress-bearing and shape-maintainingmurein sacculus of Escherichia coli. Microbiol. Mol. Biol. Rev. 62:181–203.

30. Iwai, N., T. Ebata, H. Nagura, T. Kitazume, K. Nagai, and M. Wachi. 2004.Structure-activity relationship of S-benzylisothiourea derivatives to inducespherical cells in Escherichia coli. Biosci. Biotechnol. Biochem. 68:2265–2269.

31. Iwai, N., T. Fujii, H. Nagura, M. Wachi, and T. Kitazume. 2007. Structure-activity relationship study of the bacterial actin-like protein MreB inhibitors:effects of substitution of benzyl group in S-benzylisothiourea. Biosci. Bio-technol. Biochem. 71:246–248.

32. Iwai, N., K. Nagai, and M. Wachi. 2002. Novel S-benzylisothiourea com-pound that induces spherical cells in Escherichia coli probably by acting on arod-shape-determining protein(s) other than penicillin-binding protein 2.Biosci. Biotechnol. Biochem. 66:2658–2662.

33. Iwaya, M., R. Goldman, D. J. Tipper, B. Feingold, and J. L. Strominger.1978. Morphology of an Escherichia coli mutant with a temperature-depen-dent round cell shape. J. Bacteriol. 136:1143–1158.

34. Jones, L. J., R. Carballido-Lopez, and J. Errington. 2001. Control of cellshape in bacteria: helical, actin-like filaments in Bacillus subtilis. Cell 104:913–922.

35. Karczmarek, A., R. Martinez-Arteaga, S. Alexeeva, F. G. Hansen, M. Vi-cente, N. Nanninga, and T. den Blaauwen. 2007. DNA and origin regionsegregation are not affected by the transition from rod to sphere afterinhibition of Escherichia coli MreB by A22. Mol. Microbiol. 65:51–63.

36. Kawai, Y., R. A. Daniel, and J. Errington. 2009. Regulation of cell wallmorphogenesis in Bacillus subtilis by recruitment of PBP1 to the MreB helix.Mol. Microbiol. 71:1131–1144.

37. Kruse, T., J. Moller-Jensen, A. Lobner-Olesen, and K. Gerdes. 2003. Dys-functional MreB inhibits chromosome segregation in Escherichia coli.EMBO J. 22:5283–5292.

38. Magnet, S., L. Dubost, A. Marie, M. Arthur, and L. Gutmann. 2008. Iden-tification of the L, D-transpeptidases for peptidoglycan cross-linking in Esch-erichia coli. J. Bacteriol. 190:4782–4785.

39. Markiewicz, Z., B. Glauner, and U. Schwarz. 1983. Murein structure andlack of DD- and LD-carboxypeptidase activities in Caulobacter crescentus. J.Bacteriol. 156:649–655.

40. Mashburn-Warren, L. M., and M. Whiteley. 2006. Special delivery: vesicletrafficking in prokaryotes. Mol. Microbiol. 61:839–846.

41. Noguchi, N., K. Yanagimoto, H. Nakaminami, M. Wakabayashi, N. Iwai, M.Wachi, and M. Sasatsu. 2008. Anti-infectious effect of S-benzylisothioureacompound A22, which inhibits the actin-like protein, MreB, in Shigella flex-neri. Biol. Pharm. Bull. 31:1327–1332.

42. Poindexter, J. L. S., and G. Cohen-Bazire. 1964. The fine structure of stalkedbacteria belonging to the family Caulobacteraceae. J. Cell Biol. 23:587–607.

43. Poindexter, J. S., and J. G. Hagenzieker. 1982. Novel peptidoglycans inCaulobacter and Asticcacaulis spp. J. Bacteriol. 150:332–347.

44. Priyadarshini, R., M. A. de Pedro, and K. D. Young. 2007. Role of pepti-doglycan amidases in the development and morphology of the division sep-tum in Escherichia coli. J. Bacteriol. 189:5334–5347.

45. Robertson, G. T., T. B. Doyle, Q. Du, L. Duncan, K. E. Mdluli, and A. S.Lynch. 2007. A novel indole compound that inhibits Pseudomonas aeruginosagrowth by targeting MreB is a substrate for MexAB-OprM. J. Bacteriol.189:6870–6881.

46. Rodriguez, A. D., M. J. Lear, and J. J. La Clair. 2008. Identification of thebinding of sceptrin to MreB via a bidirectional affinity protocol. J. Am.Chem. Soc. 130:7256–7258.

47. Sauvage, E., F. Kerff, M. Terrak, J. A. Ayala, and P. Charlier. 2008. Thepenicillin-binding proteins: structure and role in peptidoglycan biosynthesis.FEMS Microbiol. Rev. 32:234–258.

VOL. 192, 2010 ROD SHAPE RECOVERY IN C. CRESCENTUS 1683

Dow

nloa

ded

from

http

s://j

ourn

als.

asm

.org

/jour

nal/j

b on

14

Janu

ary

2022

by

120.

221.

189.

85.

48. Shiomi, D., M. Sakai, and H. Niki. 2008. Determination of bacterial rodshape by a novel cytoskeletal membrane protein. EMBO J. 27:3081–3091.

49. Simon, R., U. Priefer, and A. Puhler. 1983. A broad host range mobilizationsystem for in vivo genetic engineering-transposon mutagenesis in Gram-negative bacteria. Biotechnology 1:784–791.

50. Slovak, P. M., G. H. Wadhams, and J. P. Armitage. 2005. Localization ofMreB in Rhodobacter sphaeroides under conditions causing changes in cellshape and membrane structure. J. Bacteriol. 187:54–64.

51. Smit, J., D. A. Grano, R. M. Glaeser, and N. Agabian. 1981. Periodic surfacearray in Caulobacter crescentus: fine structure and chemical analysis. J. Bac-teriol. 146:1135–1150.

52. Srivastava, P., G. Demarre, T. S. Karpova, J. McNally, and D. K. Chattoraj.2007. Changes in nucleoid morphology and origin localization upon inhibi-tion or alteration of the actin homolog, MreB, of Vibrio cholerae. J. Bacteriol.189:7450–7463.

53. Thanbichler, M., A. A. Iniesta, and L. Shapiro. 2007. A comprehensive set ofplasmids for vanillate- and xylose-inducible gene expression in Caulobactercrescentus. Nucleic Acids Res. 35:e137.

54. Thanbichler, M., and L. Shapiro. 2006. MipZ, a spatial regulator coordinat-ing chromosome segregation with cell division in Caulobacter. Cell 126:147–162.

55. Uehara, T., and J. T. Park. 2008. Growth of Escherichia coli: significance ofpeptidoglycan degradation during elongation and septation. J. Bacteriol.190:3914–3922.

56. van den Ent, F., L. A. Amos, and J. Lowe. 2001. Prokaryotic origin of theactin cytoskeleton. Nature 413:39–44.

57. Varma, A., M. A. de Pedro, and K. D. Young. 2007. FtsZ directs a secondmode of peptidoglycan synthesis in Escherichia coli. J. Bacteriol. 189:5692–5704.

58. Vats, P., and L. Rothfield. 2007. Duplication and segregation of the actin(MreB) cytoskeleton during the prokaryotic cell cycle. Proc. Natl. Acad. Sci.U. S. A. 104:17795–17800.

59. Vollmer, W., and U. Bertsche. 2008. Murein (peptidoglycan) structure,architecture and biosynthesis in Escherichia coli. Biochim. Biophys. Acta1778:1714–1734.

60. Vollmer, W., D. Blanot, and M. A. de Pedro. 2008. Peptidoglycan structureand architecture. FEMS Microbiol. Rev. 32:149–167.

61. Vollmer, W., and J. V. Holtje. 2004. The architecture of the murein (pepti-doglycan) in gram-negative bacteria: vertical scaffold or horizontal layer(s)?J. Bacteriol. 186:5978–5987.

62. Wachi, M., M. Doi, Y. Okada, and M. Matsuhashi. 1989. New mre genesmreC and mreD, responsible for formation of the rod shape of Escherichiacoli cells. J. Bacteriol. 171:6511–6516.

63. Wachi, M., and M. Matsuhashi. 1989. Negative control of cell division bymreB, a gene that functions in determining the rod shape of Escherichia colicells. J. Bacteriol. 171:3123–3127.

64. Wagner, J. K., C. D. Galvani, and Y. V. Brun. 2005. Caulobacter crescentusrequires RodA and MreB for stalk synthesis and prevention of ectopic poleformation. J. Bacteriol. 187:544–553.

65. Walker, S. G., D. N. Karunaratne, N. Ravenscroft, and J. Smit. 1994. Char-acterization of mutants of Caulobacter crescentus defective in surface attach-ment of the paracrystalline surface layer. J. Bacteriol. 176:6312–6323.

66. Wang, Y., B. D. Jones, and Y. V. Brun. 2001. A set of ftsZ mutants blockedat different stages of cell division in Caulobacter. Mol. Microbiol. 40:347–360.

67. Young, K. D. 2006. The selective value of bacterial shape. Microbiol. Mol.Biol. Rev. 70:660–703.

1684 TAKACS ET AL. J. BACTERIOL.

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nloa

ded

from

http

s://j

ourn

als.

asm

.org

/jour

nal/j

b on

14

Janu

ary

2022

by

120.

221.

189.

85.