Regulated proteolysis of a cross-link–specificpeptidoglycan hydrolase contributes tobacterial morphogenesisSantosh Kumar Singh1, Sadiya Parveen, L SaiSree, and Manjula Reddy2

Centre for Cellular and Molecular Biology, Hyderabad, India 500007

Edited by Joe Lutkenhaus, University of Kansas Medical Center, Kansas City, KS, and approved July 27, 2015 (received for review April 21, 2015)

Bacterial growth and morphogenesis are intimately coupled to ex-pansion of peptidoglycan (PG), an extensively cross-linked macro-molecule that forms a protective mesh-like sacculus around thecytoplasmic membrane. Growth of the PG sacculus is a dynamicevent requiring the concerted action of hydrolases that cleave thecross-links for insertion of new material and synthases that catalyzecross-link formation; however, the factors that regulate PG expan-sion during bacterial growth are poorly understood. Here, we showthat the PG hydrolase MepS (formerly Spr), which is specific to cleav-age of cross-links during PG expansion in Escherichia coli, is modulatedby proteolysis. Using combined genetic, molecular, and biochemicalapproaches, we demonstrate that MepS is rapidly degraded by aproteolytic system comprising an outer membrane lipoprotein ofunknown function, NlpI, and a periplasmic protease, Prc (or Tsp). Insummary, our results indicate that the NlpI–Prc system contributesto growth and enlargement of the PG sacculus by modulating thecellular levels of the cross-link–cleaving hydrolase MepS. Overall,this study signifies the importance of PG cross-link cleavage and itsregulation in bacterial cell wall biogenesis.

bacterial morphogenesis | peptidoglycan | regulated proteolysis | MepS |NlpI-Prc

Peptidoglycan (PG or murein) is a unique and essential con-stituent of eubacterial cell walls, thus making it an excellenttarget for several antimicrobial agents. It is a single, large, ex-tensively cross-linked macromolecule that forms a mesh-likesacculus protecting cells against intracellular turgor pressure inaddition to conferring cell shape. Structurally, the PG sacculus ismade up of linear glycan strands cross-linked to each other byshort peptide chains forming a continuous layer around the cy-toplasmic membrane. The glycan strands are made up of alter-nating N-acetyl muramic acid (NAM) and N-acetyl glucosamine(NAG) disaccharide units in which NAM is covalently attachedto a peptide chain containing 2- to 5-amino acid residues, withthe pentapeptide consisting of L-alanine (ala)−D-glutamic acid(glu)−meso-diaminopimelic acid (mDAP)−D-ala−D-ala. Normally,D-ala of one peptide chain is cross-linked to mDAP of anotherpeptide chain of an adjacent glycan strand, resulting in an ex-tensively cross-linked single- or multilayered sacculus (1).Because the murein sacculus totally encircles the cytoplasmic

membrane, growth of a cell is tightly coupled to expansion of PG.Growth of the PG sacculus is a dynamic and coordinated eventrequiring concerted action both of murein hydrolases that fa-cilitate cleavage of cross-links for the insertion of nascent mureinmaterial and of synthases that catalyze cross-link formation be-tween adjacent glycan strands (Fig. 1) (2, 3).Escherichia coli encodes multiple PG synthases that catalyze

the formation of D-ala−mDAP cross-links in the PG sacculus.The class I enzymes (PBP1a and PBP1b, encoded by mrcA andmrcB, respectively) are bifunctional and possess both glycosyltransferase (GT) and transpeptidase (TP) activities whereas classII enzymes (PBP2 and PBP3, encoded by mrdA and ftsI, re-spectively) are monofunctional and possess only the TP activity

(4, 5). The TP activity of PBP1a and PBP1b is activated, respec-tively, by their cognate lipoprotein cofactors LpoA and LpoB,located in the outer membrane (6, 7).Given that the interconnecting peptide bridges in the PG sac-

culus need to be cleaved for the insertion of new murein material,hydrolytic enzymes with such activity are expected to be critical forPG expansion and thus for bacterial viability (2, 3, 8). Essentialcross-link–specific hydrolases have recently been identified in bothGram-positive and -negative bacteria (9–13). E. coli possessesseveral hydrolytic enzymes specific for D-ala−mDAP cross-links(termed D,D-endopeptidases based on their specificity) (14); ofthese peptidases, Spr, YebA, and YdhO (renamed MepS, MepM,and MepH, respectively, in which mep stands for murein endo-peptidase) are critical for growth under normal physiological con-ditions because a mutant lacking all these three endopeptidases isunable to incorporate new murein and exhibits rapid lysis (9).Although cross-link cleavage is fundamental for PG growth, it

is obvious that such cleavage needs to be stringently regulated atthe spatiotemporal level to avoid lethal breakage and rupture ofthe PG sacculus. In addition, to maintain the continuum andintegrity of PG, cleavage must be tightly coupled to cross-linkformation, but the underlying mechanisms are not known.To understand how this potentially lethal hydrolytic activity is

controlled in the cell, here, we examined the regulation of theD,D-endopeptidase MepS. We found that MepS is highly abundantin the exponential phase of growth, with levels falling sharply atthe onset of the stationary phase. Using combined genetic, mo-lecular, and biochemical approaches, we demonstrate that MepS

Significance

Peptidoglycan (PG) is a unique and essential cross-linked, bag-like macromolecule that completely encases the cytoplasmicmembrane and confers shape and rigidity to a bacterial cell.Therefore, bacterial cell growth is tightly coupled to PG ex-pansion, requiring the coordinate activity of hydrolases thatcleave the cross-links and synthases that catalyze the cross-linkformation. This study highlights the importance of cross-linkcleavage and its regulation in PG biogenesis by demonstratingthe modulation of a cross-link–specific PG hydrolytic enzyme,MepS, by a previously unidentified degradation system consist-ing of an outer membrane lipoprotein, NlpI and a periplasmicprotease, Prc. These studies facilitate better understanding ofbacterial cell wall synthesis, which is a target of several antimi-crobial therapeutic agents.

Author contributions: S.K.S., S.P., L.S., and M.R. designed research; S.K.S., S.P., and L.S.performed research; M.R. analyzed data; and M.R. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.1Present address: Molecular Biology Unit, Institute of Medical Sciences, Banaras HinduUniversity, Varanasi, India 221005.

2To whom correspondence should be addressed. Email: manjula@ccmb.res.in.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1507760112/-/DCSupplemental.

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is degraded rapidly by a previously unknown proteolytic systemcomprising a tetratricopeptide repeat (TPR)-containing outermembrane (OM) lipoprotein, NlpI, and a periplasmic protease,Prc. In summary, we show that the NlpI–Prc system regulates PGsynthesis by altering the levels of MepS, a hydrolase that breaksthe cross-links for insertion of new murein material during growthof the PG sacculus.

ResultsGrowth Phase Specificity of PG Hydrolase MepS. MepS is an OMlipoprotein, belonging to the NlpC/P60 superfamily of pepti-dases, that contributes to the essential murein hydrolytic activityin E. coli (9). Because this observation reflected a vital role forMepS in growth and enlargement of the PG sacculus, we exam-ined the level of cellular MepS during various phases of growth, byWestern analysis using a functional C-terminal 3XFlag-taggedderivative at the native chromosomal locus (Table S1). Fig. 2shows that the level of MepS is high throughout the exponentialphase of growth, with levels falling steeply at the onset of thestationary phase, with further gradual decrease as cells progressinto the late stationary phase.

Regulation of MepS Is Dependent on NlpI and Prc. To understandthe basis of the growth-phase specificity of MepS, we identifiedmutations that altered the activity of alkaline phosphatase (PhoA)fused to the C terminus of mepS (Para::mepS-phoA) (Table S2).Two such mutations that increased the activity of MepS-PhoAwere in loci encoding NlpI, an OM lipoprotein of unknown func-tion, and Prc, a periplasmic protease (Fig. S1A). nlpI or prc deletionmutants do not grow on media of low osmotic strength in ad-dition to forming long filaments (15, 16), and absence of MepSwas able to suppress the growth and morphological defects ofboth these mutants (Fig. S1B). Suppression of nlpI or prc mutantphenotypes by mepS deletion was abolished by a plasmid-bornecopy of WT mepS, but not by a derivative carrying a mutation,C68A, in the active site of MepS (Fig. S1C). These results sug-gest that the unfettered D,D-endopeptidase activity of MepS isresponsible for the phenotypes of nlpI or prc mutants.Because the experiments above were indicative of regulation

of MepS by NlpI and/or Prc, we measured MepS levels in the

absence of NlpI or Prc. The data in Fig. 3A show that, in nlpI orprc mutants, the level of MepS is constitutively high throughoutthe growth cycle, indicating loss of regulation.To understand the basis of MepS regulation, a pulse–chase

experiment was performed after inhibition of protein synthesisby treatment with spectinomycin. Fig. 3B shows that increasedMepS in nlpI or prc mutants is a consequence of enhanced post-translational stability. The half-life of MepS was substantially in-creased from ∼1–2 min in exponentially growing WT cells to morethan 45 min in mutants lacking NlpI or Prc. Consistent with aposttranslational model of regulation, the expression of a chro-mosomal mepS-lacZ transcriptional fusion was altered neitherduring the growth cycle nor in nlpI or prc mutants (Table S3).To examine whether NlpI and Prc act in concert to regulate

MepS, the level of MepS was measured in single and doublemutants. As shown in Fig. 4A, MepS levels remain equally highin all of these mutants, suggesting that NlpI and Prc share acommon pathway in regulation of MepS. Furthermore, a modestincrease in NlpI led to a significant decrease of MepS, which inturn was dependent on the presence of functional Prc, demon-strating the combined requirement of both these factors in MepSregulation (Fig. 4A).

Prc Does Not Process NlpI. In light of an earlier report that Prcprotease processes NlpI into a mature functional derivative bycleaving the latter’s C-terminal 10- to 11-amino acid residues(17), we measured the size of NlpI in WT and prc mutant strainsby Western analysis using anti-NlpI antisera. Fig. 4B shows that thesize and level of NlpI remain unaltered in the presence or absenceof Prc. Importantly, a plasmid-borne copy of nlpI encoding apolypeptide lacking the extreme C-terminal 10 amino acids (NlpI-D284) was able to functionally complement an nlpI mutant but nota prc or prc nlpI mutant, indicating that both NlpI and Prc con-tribute independently and directly to the stability of MepS (Fig. S2).

NlpI Overexpression Phenocopies MepS Deletion. Because multiplecopies of NlpI led to lowered MepS levels, (Fig. 4A), we exam-ined the effect of increased NlpI on the growth and viability of amutant deleted for the alternative endopeptidase MepM becausea strain doubly deficient in MepS and MepM is not viable on richmedia (9). A modest increase of NlpI resulted in severe cell lysisin the mepM mutant that was suppressed by introduction ofadditional copies of MepS (Fig. 5A and Fig. S3). In WT E. coli,as well, increased NlpI conferred the phenotypes of a mepSdeletion mutant: i.e., inability to grow on media of low osmo-larity and altered cell shape from rods to fat ellipsoids (Fig. 5B).Taken together, these data indicate that increased NlpI de-creases MepS levels very effectively in vivo and mimics the MepS−

phenotype.

Fig. 1. Schematic of PG sacculus expansion. (A) Depiction of PG sacculusenlargement during growth of a bacterial cell. New murein strands (coloredred) are incorporated into the preexisting murein strands (colored blue) toexpand the PG sacculus (2, 3). (B) A model showing cleavage and resynthesisof cross-links for successful insertion of new glycan strands for expansion ofthe PG sacculus (8). The green bars indicate peptide stems whereas the smalldark red bars represent the cross-bridges between the peptide stems ofadjacent glycan strands.

Fig. 2. Growth phase specificity of MepS. Strain MR802 (MG1655 mepS-Flag) was grown in LB and fractions were collected at various time intervals(with indicated A600 values) during the growth cycle. Normalized cell extracts(corresponding to 0.25 OD cells) were separated by SDS/PAGE, and MepS wasdetected by Western analysis using anti-Flag antisera. FtsZ was used as aloading control.

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NlpI Enhances the Proteolysis of MepS by Prc. Our results thus farsuggested that NlpI and Prc together modulate the level of MepSin vivo. To test this observation directly in vitro, we purified MepS,NlpI, and Prc, as C-terminal hexahistidine fusion proteins (Fig.6A, lanes 2, 3, and 4) lacking their signal sequences, and per-formed degradation assays (Fig. 6A). NlpI and MepS were stablein the presence of one another (Fig. 6A, lane 5) as was Prc in thepresence of NlpI (Fig. 6A, lane 6). Coincubation of Prc andMepS led to degradation of the latter to a minimal extent (Fig.6A, compare lanes 7 and 8), and this process was substantiallyaccelerated in the presence of NlpI (Fig. 6A, compare lanes 9 and10). A time course experiment also indicated that MepS degra-dation by Prc is enhanced to a large extent by addition of NlpI(Fig. S4 A and B). Degradation of MepS by Prc was abolished byintroduction of mutations at the active site residues of Prc (S452Aor K477A) (18), even in the presence of NlpI (Fig. 6B, comparelanes 5, 6, and 7). The proteolytic activity of Prc toward MepS wasspecific because two other putative periplasmic peptidases, NlpC(a paralog of MepS belonging to the NlpC/P60 family) and YfiH,were not considerably degraded by Prc (Fig. S4C). Overall, theseresults demonstrate that MepS is a substrate of Prc protease andthat NlpI facilitates this degradation.

NlpI Facilitates Interaction of MepS and Prc in Vivo. To examine thein vivo interactions of MepS with Prc and/or NlpI, pull-downassays were performed in strains carrying a functional chro-mosomal Prc-HA fusion allele, along with a plasmid-encodedfunctional MepS bearing a hexahistidine tag at its C terminus(Para::mepS-His). Immunoblot analysis of the MepS-His pull-down fractions showed that NlpI, but not Prc, is copurified withMepS (Fig. 7A and Fig. S5A). The MepS–NlpI interaction oc-curred even in absence of Prc (Fig. 7A). A mutant derivative ofPrc-HA carrying an active site mutation, S452A, that is expectedto bind but not cleave MepS (Fig. S5B) also could not be cop-urified with MepS, suggesting that Prc does not directly interactwith MepS in vivo (Fig. 7A).Pull-down assays were also done using a plasmid-borne NlpI-

His (Para::nlpI-His) in strains carrying functional MepS-Flag andPrc-HA fusion alleles at their native chromosomal loci. Here,

both MepS and Prc were copurified with NlpI (Fig. 7B), andthese interactions were independent of Prc and MepS, respectively(Fig. 7B). Taken together, these results indicate that NlpI is able toform binary complexes each with MepS and Prc in vivo, but MepSis itself unable to form a binary complex with Prc, suggesting thatNlpI is required as an adapter protein to bring together the Prcprotease with its target protein MepS, an interpretation that is con-sistent with the genetic and biochemical data.

DiscussionIn this study, we demonstrate that a previously unknown pro-teolytic system comprising NlpI and Prc contributes to growth ofthe PG sacculus by rapidly degrading the cross-link–specific keymurein hydrolytic enzyme MepS. This stringent regulation sug-gests that the cross-link cleavage activity of MepS is a crucialdeterminant of PG synthesis and may indeed be the rate-limitingstep in PG expansion. To our knowledge, MepS is the first ex-ample of an enzyme involved in PG metabolism to be regulatedby proteolysis.The mechanism of degradation of MepS by the NlpI–Prc

system seems to be analogous to that described earlier on theproteolysis of the rate-limiting enzyme of the lipopolysaccharidebiosynthesis, LpxC, by an essential membrane-anchored proteaseFtsH and the TPR-containing adapter protein YciM (19).

Role of NlpI–Prc in E. coli. Contrary to an earlier report that hadsuggested a proteolytic processing role for Prc to activate NlpIinto a mature functional derivative (17), our results show thatNlpI is not processed by Prc either in vivo (Fig. 4B) or in vitro(Fig. 6A). Rather, we find that NlpI binds both Prc and MepS,

Fig. 3. Regulation of MepS is dependent on NlpI and Prc. (A) MepS-Flaglevels in nlpI and prc mutants. Strains MR802 (MG1655 mepS-Flag), MR803(MR802 ΔnlpI), or MR804 (MR802 Δprc) were grown in LB, and fractionswere collected at different time points during growth and analyzed byWestern blotting as described in the legend to Fig. 2. (B) Determination ofstability of MepS by pulse–chase experiment (in vivo degradation assay). Torapidly growing cultures of the above strains in LB (at an OD600 of ∼0.4),spectinomycin (Spec) was added at a concentration of 300 μg/mL to blocktranslation, and fractions were collected at indicated time points and ana-lyzed as described in the legend to Fig. 2.

Fig. 4. Requirement of both NlpI and Prc for regulation of MepS. (A) Levelsof MepS in single and double mutants of nlpI and prc. Strains MR802(MG1655 mepS-Flag), MR803 (MR802 ΔnlpI), MR804 (MR802 Δprc), andMR806 (MR802 ΔnlpI Δprc) were grown in LB to an A600 of 1.0, and MepSwas detected by Western analysis as described in the legend to Fig. 2. Plas-mid-carrying strains were grown with appropriate antibiotic (Spec) and1 mM IPTG. FtsZ was used as a loading control. (B) NlpI is not processed byPrc. The indicated strains MG1655 (WT), MR810 (MG1655 ΔmepS), MR812(MG1655 Δprc), MR814 (MG1655 ΔmepS Δprc) were grown in LB and har-vested at an A600 of ∼1.0. Normalized cell extracts were analyzed by Westernblotting using anti-NlpI antisera. Plasmid-carrying strains were grown withSpec and 1 mM IPTG. The C-terminal truncations of NlpI (bands in lanes 8 and9) served as controls to indicate the size of processed NlpI. Processed NlpI isexpected to be of 284 amino acids in length (17) whereas full-length is 294amino acids.

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thereby acting like an adapter protein to facilitate degradation ofMepS by Prc (Figs. 6 and 7). NlpI is an OM lipoprotein withmultiple TPR motifs (which is conserved in γ-proteobacteria ofthe Enterobacteriaceae, Pasteurellaceae, and Vibrionaceae line-ages) (20). It is known that TPR proteins mediate protein–protein

interactions to facilitate assembly of multiprotein complexes (21).NlpI binding to Prc has been shown earlier (17). FtsI, the essentialdivision-specific transpeptidase, is also shown to be processed byPrc although the physiological significance of such processing isnot clear (22). A previous report had indicated that Prc is a per-iplasmic protein associated with the cytoplasmic membrane (16);however, as predicted by several algorithms, we find that Prc is asoluble periplasmic protein (Fig. S6). In this context, it is to benoted that Prc is specific to proteins with nonpolar C termini andthus is also called a Tail-specific protease (Tsp) (23). However, inmany of our in vitro and in vivo experiments, we have used MepSderivatives with various C-terminal tags; therefore, it is possiblethat the extent of MepS degradation observed here is an un-derestimate and that the native cellular MepS in vivo may perhapsbe more susceptible to regulation by the NlpI–Prc system.Regulation of MepS seems to be the primary role of NlpI and

Prc because most NlpI− or Prc− phenotypes are abrogated by theabsence of MepS (this study and refs. 24–27). It is to be notedthat the levels of both NlpI and Prc are constitutive and notdependent on growth cycle (Fig. S7). Interestingly, mepS (for-merly, spr) was identified initially as a suppressor of prc (24). Ourresults provide a mechanistic basis for earlier observations fromseveral groups that absence of MepS suppresses various pheno-types of prc and nlpI mutants (24–27).

Models for Regulation of MepS. The data from Fig. 2 show that thelevels of MepS significantly decline at the onset of the stationaryphase. At the same time, the pulse–chase experiments (Fig. 3B)indicate that MepS has a short half-life of ∼1–2 min, even inexponentially growing cultures, suggesting a rapid and constitu-tive degradation of MepS throughout the growth cycle. Based onthe above findings, the following two scenarios for degradationof MepS can be envisaged. In the first case, the synthesis ofMepS could be reduced in the stationary phase, therefore lead-ing to the observed decline in steady-state levels as it is rapidlydegraded. However, then the question arises about why cellsshould indulge in a futile cycle of simultaneous synthesis anddegradation unless such a mechanism has evolved to permit in-stantaneous changes in the levels of MepS in individual cellswithin a population. The second scenario would be that the ob-served half-life of MepS represents an average of different cell

Fig. 5. NlpI overexpression phenocopies MepS− phenotype. (A) Lethal ef-fect of overexpressed NlpI in ΔmepM mutants. Strains MR816 (BW27783ΔmepM) carrying either pTRC99a (Ptrc) or pMN204 (Ptrc::nlpI) were grown inLB with Amp, and 5 μL of various dilutions were placed on indicated platesand grown. IPTG was used at 100 μM. (B) Effect of overexpressed NlpI in WTE. coli. MG1655 carrying either pTRC99a (Ptrc) or pMN204 (Ptrc::nlpI) wascultured and grown on nutrient agar (NA) at 42 °C. IPTG was used at 100 μM.ΔmepS strains are known not to grow on NA at 42 °C (24). For microscopy,the cultures were grown, diluted 1:100 either in LBON (for WT and ΔmepSstrains) or in LBON plus 100 μM IPTG (for plasmid-carrying strains) and grownat 42 °C until an OD600 of ∼0.4–0.6. Differential interference contrast (DIC)images were taken after concentrating and spotting the culture on agarosepads. Overexpression of NlpI in WT is known to result in loss of rod shapeand formation of prolate ellipsoid cells (15). It is not clear why the mutantslacking MepS (or those overproducing NlpI) exhibit fat and wide cell mor-phology; these cells may possibly be partially deficient in cross-link cleavage,leading to defective growth and elongation of the PG sacculus.

Fig. 6. MepS is a substrate of Prc. (A) In vitro degradation assays. The C-terminal hexahistidine-tagged MepS, NlpI, and Prc proteins were mixed in allcombinations (as indicated) and incubated for 3 h at 37 °C followed by SDS/PAGE and Coomassie blue staining. Each reaction contained the following(approximately): MepS, 15 μg; NlpI, 2 μg; and Prc, 4 μg. The mixtures in lanes 7 and 9 served as controls (0 h; no incubation). M is a protein marker with thefollowing molecular size range (in kDa): 170, 130, 100, 70, 55, 40, 35, 25, 15, and 10. Purified MepS was always seen on SDS/PAGE as a doublet, and N-terminalsequencing confirmed that both bands belong to MepS itself. (B) Specificity of Prc. Two mutant derivatives of Prc (carrying K477A or S452A alterations at theactive site residues) did not show any proteolytic activity (even when higher concentrations were used) against MepS in the presence of NlpI.

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cycle-specific values for the asynchronously growing culture. Inthis model, MepS is relatively stable in cells that are in theelongation mode of PG synthesis compared with the cells thatare in the division mode of PG synthesis (that is, when the re-quirement for expansion of the PG sacculus is low). The abovetwo possibilities may not be mutually exclusive and need to beexperimentally tested further.However, additional questions that remain to be addressed

relate to the nature of the signal generated during expansion ofthe PG sacculus and how it is transmitted to NlpI, an OM li-poprotein. The causes of signal generation may include inputsfrom the PG biosynthetic machinery, alterations in membraneturgor, or the process of cross-link formation itself. Other instancesof regulation of PG synthesis by OM lipoproteins are known: TheTP activity of PG synthases PBP1a and PBP1b is activated bybinding of OM lipoproteins LpoA and LpoB, respectively (6, 7).How these OM lipoproteins sense, bind, and regulate their cognateeffectors is an interesting question to be addressed.

Regulation of Other PG Hydrolases. Most eubacterial genomes en-code a multitude of PG hydrolases that function in murein growth,maturation, turnover, recycling, autolysis, and cleavage of septum

at cell division (12–14). Even as murein hydrolytic activity is es-sential for bacterial growth and division, it needs also to be tightlyregulated to prevent otherwise lethal degradation of PG. Acti-vation of division-specific amidases in E. coli has been shown tobe coupled to formation of a cytokinetic ring at the midcell inwhich a divisomal component, FtsEX, activates EnvC, a catalyti-cally inactive LytM family peptidase that in turn activates theamidases to cleave septal PG for separation of daughter cells (28).In Bacillus subtilis, two L,D-endopeptidases, CwlO and LytE,

form a minimal essential set of hydrolases for PG enlargement(10, 12, 13), of which CwlO is activated by the FtsEX complex(29) and LytE is regulated by MreBH (30). Likewise, FtsEXactivates the hydrolytic activity of the peptidases PcsB and RipCin Streptococcus pneumoniae and Mycobacterium tuberculosis, re-spectively (31, 32). In contrast, here, we find that MepS is regu-lated by proteolysis, possibly providing an advantage of a rapidresponse to changing environmental conditions.

Coupling of Cross-Link Cleavage and Cross-Link Formation. For aneffective enlargement of the PG sacculus, hydrolysis is as im-portant as synthesis although the coupling between these pro-cesses is not well understood. In Gram-negative organisms withmonolayered PG sacculus, cleavage and resynthesis are believedto be either concomitant or tightly coupled to maintain PG in-tegrity (2, 3, 33–36). On the other hand, in Gram-positive organ-isms such as B. subtilis, in which the PG sacculus is multilayered,hydrolysis is not coupled to synthesis even though PG hydrolyticactivity is essential for growth (10, 12, 13, 29).It is not yet clear whether cross-link cleavage initiates or fol-

lows the cross-link formation in E. coli. However, because someof our preliminary results suggest additional roles for NlpI and/orPrc in PG cross-link formation, we believe that the NlpI–Prc systemmay facilitate concomitant cleavage and resynthesis of cross-links, resulting in successful expansion of the PG sacculus duringgrowth of bacteria.

Materials and MethodsDetailed strain and plasmid constructions, additional materials, methods,Tables S1–S3, and Figs. S1–S7 are listed in SI Materials and Methods.

Media and Growth Conditions. Strains were normally grown in LB (1% tryp-tone, 0.5% yeast extract, 1% NaCl) unless otherwise indicated. LBON has noadded NaCl. Nutrient broth has 0.5% peptone and 0.3% beef extract. Solidmedia had agar to a concentration of 1.5% (wt/vol). Antibiotics were used atthe following concentrations (μg·mL−1): ampicillin (Amp), 50 μg·mL−1; chlor-amphenicol (Cam), 25 μg·mL−1; kanamycin (Kan), 25 μg·mL−1; and spectino-mycin (Spec), 50 μg·mL−1. Concentrations of L-arabinose and isopropyl β-D-thiogalactopyranoside (IPTG) were indicated. Growth temperature was 37 °Cunless otherwise specified.

Western Blotting. Samples were separated using SDS/PAGE and transferredonto a nitrocellulose or PVDF membrane. Membrane was blocked by 5%skimmed milk in 1× TBS-T (Tris-NaCl-Tween 20) for 2 h and then incubatedovernight with primary antibodies (1:10,000 for α-NlpI, 1:3,000 for α-His,α-FLAG, and α-HA, and 1:5,000 for α-FtsZ) at 4 °C. Membrane was washedthree times with 1× TBS-T and then probed with secondary antibodies(1:10,000) tagged with horseradish peroxidase (HRP) and incubated for 1 hat room temperature. Membrane was overlaid with ECL Prime detectionsubstrate (Amersham) for 5 min, and the blots were developed.

Pull-Down Experiments. Pull downs were performed as described earlier, butwith modifications (6). Strains were grown overnight in LB broth supple-mented with Amp and next morning diluted 1:100 into fresh LB (200 mL)and grown at 37 °C until an OD600 of ∼0.2. At this point, 0.05% arabinose wasadded, and growth was continued until OD600 was 0.8–1.0. Cells were re-covered by centrifugation, and the cell pellet was washed, resuspended in25 mL of the lysis buffer (50 mM Tris·Cl, 10 mM MgCl2, 100 mM NaCl, 20%glycerol, 2% Triton X-100, 1 mg/mL lysozyme, 1× mixture protease inhibitor,20 units of DNase, 20 units of RNase, and 10 mM imidazole; pH 8.0), andincubated on ice for 2 h. This mixture was subjected to brief sonication, andthe lysate was gently stirred overnight with glass beads at 4 °C to solubilize

Fig. 7. Interactions of MepS, NlpI, and Prc. (A) Western blot showing in-teraction of MepS-His with NlpI and Prc. Strains MR818 (MG1655 ΔmepS prc-HA-Cam), MR819 (MG1655 ΔmepS prcS452A-HA-Cam), or MR814 (MG1655ΔmepS Δprc) carrying pMN217 (Para::mepS-His) were grown overnight inLB with Amp and next morning diluted 1:100 into fresh LB and at A600 of∼0.2–0.3 expression of MepS was induced by addition of 0.05% L-arabinose.Cultures were further grown until an A600 of ∼0.8–1.0, and MepS-His waspurified using Ni2+-NTA beads as described in SI Materials and Methods. Thepurified fractions of MepS-His (pull-down fractions, PL) along with the inputsamples (IN) were separated on SDS/PAGE, and immunoblot analysis wasperformed to detect MepS, NlpI, and Prc with anti-His, anti-NlpI, and anti-HAantibodies, respectively. All strains had a deletion of mepS to reduce theinterference from the endogenous MepS protein. A strain carrying plasmid-borne, untagged MepS (MG1655 ΔmepS/Para::mepS) was used as a negativecontrol (Fig. S5A). (B) Western blot showing interaction of NlpI-His withMepS and Prc. Strains MR805 (MG1655 ΔnlpI mepS-Flag), MR806 (MG1655ΔnlpI Δprc mepS-Flag), MR820 (MG1655 ΔnlpI mepS-Flag prc-HA-Cam),MR821 (MG1655 ΔnlpI ΔmepS prc-HA-Cam), and MR822 (MG1655 ΔnlpI prc-HA-Cam) carrying pMN218 (Para::nlpI-His) were grown with 0.05% L-arabi-nose to an OD of 0.8–1.0. NlpI-His was purified from all these strains using anNi2+-NTA column, and the purified fractions (PL) along with the input frac-tions (IN) were separated on SDS/PAGE. Western analysis was done usinganti-His, anti-Flag, and anti-HA antibodies to detect NlpI, MepS, and Prc,respectively. In both these experiments, input sample corresponds to ∼0.3 ODcells whereas the pull-down fraction is ∼20-fold concentrated (i.e., from∼6 OD cells).

10960 | www.pnas.org/cgi/doi/10.1073/pnas.1507760112 Singh et al.

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the membrane proteins. The next morning, insoluble material of the lysatewas removed by centrifugation (14,000 × g, 15 min, 4 °C), at which point100 μL was set aside for input fractions, and then the supernatant was mixedwith 200 μL of Ni2+-NTA agarose and mixed for 2 h at 4 °C. The agarosebeads were washed twice with 10 mL of wash buffer (50 mM Tris·Cl, 100 mMNaCl, and 20 mM imidazole; pH 8.0) and twice with wash buffer containing50 mM imidazole, and the bound proteins were eluted with 250 μL of theelution buffer (50 mM Tris·Cl, 100 mM NaCl, and 300 mM imidazole; pH 8.0).

The eluate was electrophoresed using 12% SDS/PAGE, and the proteins weredetected by Western blotting.

ACKNOWLEDGMENTS. We thank Sujata Kumari and M. R. Sunayana forplasmid constructions; Nilanjan Som for Prc localization studies; Tom Bernhardtfor FtsZ antisera; and J. Gowrishankar and N. Madhusudhana Rao for advice.This work is supported by the Department of Biotechnology and the Council ofScientific and Industrial Research, Government of India. S.P. acknowledgesfinancial support from the University Grants Commission of India.

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