what does cell wall synthesis mean

Microbe Notes

Cell Wall Synthesis Inhibitors: Examples, Inhibition, Resistance

Generally, the bacterial cell consists of a cell wall, cell membrane, and nucleoid.  The cell wall is the outer covering of the bacteria-containing peptidoglycan layer which is made up of cross-linked polymers. Peptidoglycan is mainly responsible for all the mechanisms including resistivity, and virulence factors including- the shape of the bacteria.

Table of Contents

Interesting Science Videos

What are Cell wall synthesis inhibitors?

Cell wall synthesis inhibitors include antibiotics of class β- lactams and Glycopeptides. Β- lactam antibiotics consist of Penicillins, Cephalosporins, Monobactams, and Carbapenem. Glycopeptides include Vancomycin and Teicoplanin which are the most commonly used antibiotics of this class. Penicillin includes Ampicillin, Oxacillin, Cephalosporins include Cefpodoxime, Cefepime, Monobactams include Aztreonam which is the only commercially available antimicrobial of this class.  Carbapenem consists of Imipenem, Meropenem and Ertapenem. Cell wall synthesis inhibitors are the most used antibiotics for treating Gram-negative as well as Gram-positive infections.

Table: Cell wall biosynthesis inhibitors and their targets. Table Source: https://doi.org/10.1039/C6MD00585C

• Peptidoglycan is made up of a polymer of peptides and glycan. Mainly synthesis of peptidoglycan occurs in three stages which include; cytoplasm, membrane, and periplasm.
• In the cytoplasmic stage, UDP-GLcNAc is catalyzed in four steps process. MurA and MurB catalyze forming UDP-MurNAc. Four enzymes Mur C, Mur D, Mur E, and Mur F catalyzes alanine, glutamic acid which are involved in important steps. Alanine ligase and Alanine racemase are two important enzymes involved in the formation of D-Ala-D-Ala.
• The second membrane-associated stage includes the formation of lipid intermediates.
• In the third stage, peptidoglycan chains were formed by glycosyltransferases which are then cross-linked by the enzyme transpeptidases. This enzyme transpeptidase consists of  PBPs (Penicillin Binding Proteins) and Rod A
• Mur enzymes, Transpeptidases, glycosylases, and lipid components are the main components involved in cell wall synthesis.

Examples of Cell wall synthesis inhibitors

Beta-lactam antibiotics.

a. Penicillins- Ampicillin, Amoxicillin, Piperacillin, Oxacillin

b. Cephalosporins: 

• First-generation: Cefazolin, Cefalexin
• Second-generation: Cefoxitin, Cefuroxime
• Third-generation: Cefixime, Ceftazidime, Ceftriaxione
• Fourth-generation: Cefepime

c. Monobactams: Aztreonam, the only commercially available monobactam.

d. Carbapenems: Imipenem, Meropenem

Glycopeptides

Vancomycin, Oritavancin, Teicoplanin, Telavancin, Bleomycin, Ramoplanin, Decaplanin.

Process/Steps of Inhibition

• Beta-lactams interfere in synthesis by acting as a component of D-Alanine- D- Alanine with the help of transpeptidase enzyme in transpeptidase reaction. Glycine residues cross-link amino acid portion of peptide-chain in the presence of penicillin-binding proteins (PBPs). New peptidoglycan chains are formed resulting in cell rupture due to osmotic lysis. 
• Bacterial cells contain autolytic enzymes which are known for cell growth and division and beta-lactam antibiotics affect the autolytic activity resulting in bacterial cell death.
• Glycopeptide antibiotics bind to lipid  II component that prevents the recycling of central lipid transporter that plays important role in the mode of action. They inhibit peptidoglycan synthesis by linking with the pentapeptide chains and prevent the addition of new peptidoglycan units. Transglycosylation and transpeptidation are inhibited by this interaction resulting in bacterial cell death.
• Vancomycin, a last resort drug of glycopeptides, prevents the binding of D-alanyl with penicillin-binding proteins which inhibits cell wall synthesis causing bacterial lysis.
• Permeabilization and depolarization of bacterial cell membrane resulting in inhibition of cell wall synthesis by some of the glycopeptides.

Mechanisms of Resistance

 The affinity of the drug, its permeability, and stability determines the activity against bacteria. During the past two decades, resistant organisms towards antibiotics are increasing which is of great concern. Organisms develop resistance to beta-lactam antibiotics through a synthesis of PBP2a (penicillin-binding protein 2a). It has a low affinity allowing transpeptidase activity which results in bacterial colonization in patients. Chromosomal mutations also directly or indirectly increase the level of penicillin-binding protein 2a and transcription causes bacterial resistance. Β-lactams resistance is mainly due to the alteration of Penicillin-binding proteins and enzymatic degradation. Glycopeptides resistivity is due to the alteration of targets.

• Both Gram-positive and Gram-negative organisms produce beta-lactamases.  In Gram-positive organisms, penicillinase is the enzyme causing resistance towards antibiotics and can also cause a mutation by altering the enzymatic activity. 
• Gram-negative organisms can express both chromosomal as well as plasmid-mediated enzymes that have broad activity in causing resistivity.
• Resistant to methicillin and oxacillin in S. aureus staphylococcal cassette chromosome mec which contains mec A gene. PBP2a is encoded by mec A gene which shows high resistance to beta-lactam antibiotics.
• Different antibiotics are exported from cells by membrane proteins to maintain their concentration called efflux pumps. These pumps carry a wide range of antibiotics that contribute to forming MDR organisms.
• D-alanyl-alanine is changed to D-alanyl-lactate which inhibits the cross-linking of glycopeptides hence causing resistance.
• Seven van genes are responsible for causing vancomycin resistance.  These genes encode dehydrogenases that form lactate which is important for the formation of unmodified peptidoglycan.
• Characterization of different enzymes that are required for the transfer of plasmid results in vancomycin resistance.

• Canzani D and Aldeek, F. (2017). Penicillin G ‘s function, metabolites, allergy, and resistance. 1(1).
• Ghooi, R. B  and  Services SC. (2018). Inhibition of Cell Wall Synthesis -Is this the Mechanism of Action of Penicillins ? 9877(April), 2–7. https://doi.org/10.1016/0306-9877(95)90085-3
• Kang H and Park Y (2015). Glycopeptide Antibiotics : Structure and Mechanisms of Action. 45(2), 67–78.
• Kapoor G, Saigal S and Elongavan A (2017). Action and resistance mechanisms of antibiotics: A guide for clinicians.  Journal of Anaesthesiology and clinical pharmacology. 33: 300-305
• Nikolaidis I and Dessen A (2014). Resistance to antibiotics targeted to the bacterial cell wall. 23, 243–259. https://doi.org/10.1002/pro.2414
• Scheffers, D and Pinho M G. (2005). Bacterial Cell Wall Synthesis : New Insights from Localization Studies. 69(4). 585–607. https://doi.org/10.1128/MMBR.69.4.585
• Singh M, Chang J, Coffman L, Kim S J and States U. (2018). HHS Public Acces s . 121(16), 3925–3932. https://doi.org/10.1021/acs.jpcb.7b00324.A
• https://www.researchgate.net/publication/268793620_Multidrug_efflux_pumps_in_Gramnegative_ bacteria_and_their_role_in_antibiotic_resistance- 6%
• https://www.sciencedirect.com/topics/neuroscience/peptidoglycan- 4%
• https://www.sciencedirect.com/topics/immunology-and-microbiology/peptidoglycan- 3%
• https://www.sciencedirect.com/topics/biochemistry-genetics-and-molecular-biology/peptidoglycan- 2%
• https://www.sciencedirect.com/science/article/abs/pii/S0045206814000224- 1%
• https://globalrph.com/infectious-disease-list/monobactams/- 1%
• https://pubs.acs.org/doi/10.1021/acs.jpcb.7b00324- 1%
• https://pubs.asahq.org/anesthesiology/article/129/2/335/17980/Presumed-Lactam-Allergy-and-Crossreactivity- in- 1%
• https://docs.ufpr.br/~microgeral/Regulacaopeptideoglicana.pdf- 1%

About Author

Reeju Sharma

2 thoughts on “Cell Wall Synthesis Inhibitors: Examples, Inhibition, Resistance”

Bacterial cells do not have a nucleus.

Thank you, we mean to say Nucleoid.

Leave a Comment Cancel reply

Save my name, email, and website in this browser for the next time I comment.

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

• View all journals
• Explore content
• About the journal
• Publish with us
• Sign up for alerts
• Open access
• Published: 12 September 2022

Cell wall synthesis and remodelling dynamics determine division site architecture and cell shape in Escherichia coli

• Paula P. Navarro   ORCID: orcid.org/0000-0002-9123-1132 1 , 2   na1 ,
• Andrea Vettiger 3   na1 ,
• Virly Y. Ananda 1 ,
• Paula Montero Llopis 4 ,
• Christoph Allolio   ORCID: orcid.org/0000-0002-7525-9266 5 ,
• Thomas G. Bernhardt   ORCID: orcid.org/0000-0003-3566-7756 3 , 6 &
• Luke H. Chao   ORCID: orcid.org/0000-0002-4849-4148 1 , 2  

Nature Microbiology volume  7 ,  pages 1621–1634 ( 2022 ) Cite this article

18k Accesses

29 Citations

76 Altmetric

Metrics details

• Bacterial development
• Bacterial evolution
• Cell growth
• Cellular microbiology
• Cryoelectron tomography

The bacterial division apparatus catalyses the synthesis and remodelling of septal peptidoglycan (sPG) to build the cell wall layer that fortifies the daughter cell poles. Understanding of this essential process has been limited by the lack of native three-dimensional views of developing septa. Here, we apply state-of-the-art cryogenic electron tomography (cryo-ET) and fluorescence microscopy to visualize the division site architecture and sPG biogenesis dynamics of the Gram-negative bacterium Escherichia coli . We identify a wedge-like sPG structure that fortifies the ingrowing septum. Experiments with strains defective in sPG biogenesis revealed that the septal architecture and mode of division can be modified to more closely resemble that of other Gram-negative ( Caulobacter crescentus ) or Gram-positive ( Staphylococcus aureus ) bacteria, suggesting that a conserved mechanism underlies the formation of different septal morphologies. Finally, analysis of mutants impaired in amidase activation (Δ envC Δ nlpD ) showed that cell wall remodelling affects the placement and stability of the cytokinetic ring. Taken together, our results support a model in which competition between the cell elongation and division machineries determines the shape of cell constrictions and the poles they form. They also highlight how the activity of the division system can be modulated to help generate the diverse array of shapes observed in the bacterial domain.

Similar content being viewed by others

A two-track model for the spatiotemporal coordination of bacterial septal cell wall synthesis revealed by single-molecule imaging of FtsW

FtsN maintains active septal cell wall synthesis by forming a processive complex with the septum-specific peptidoglycan synthases in E. coli

Peptidoglycan synthesis drives a single population of septal cell wall synthases during division in Bacillus subtilis

Bacterial cells are typically surrounded by a multi-layered cell envelope of varying complexity depending on species 1 . Gram-positive bacteria possess a single membrane surrounded by a thick cell wall, whereas Gram-negative bacteria have a thinner wall covered by an outer membrane (OM) 2 . The cell wall determines cell shape and protects cells against osmotic lysis 3 . It is assembled from peptidoglycan (PG), which consists of glycan chains with repeating disaccharide units of N -acetylglucosamine (GlcNAc) and N -acetylmuramic acid (MurNAc). Short peptides are attached to each MurNAc sugar and used to form amide crosslinks between adjacent glycans, generating a covalent mesh encapsulating the cytoplasmic membrane.

Rod-shaped cells such as Escherichia coli ( E. coli ) lengthen their cell body through the action of the elongation machinery (Rod complex, elongasome), which incorporates new PG material at dispersed locations throughout the cylinder 3 . Cell division is then initiated by the tubulin-like FtsZ protein, which at midcell forms the Z-ring that recruits dozens of proteins to the division site, assembling the divisome 4 , 5 . This apparatus promotes localized synthesis of PG to generate the cross-wall/septum that divides the daughter cell compartments 3 . The septal PG (sPG) produced initially connects the daughters such that it must be processed to separate the newly formed cells 3 .

Our understanding of cell envelope biogenesis during cell division has been greatly influenced by electron micrographs of developing septa 6 . However, sPG has not been clearly visualized in the septa of Gram-negative bacteria. Furthermore, whether the different septal architectures observed in diverse bacteria 7 , 8 , 9 , 10 , 11 , 12 reflect fundamental differences in the division mechanism between species or arise from changes in the spatiotemporal regulation of conserved processes remains a major outstanding question. We therefore investigated the structure and dynamics of the septal PG layer of E. coli using both in situ cryo-electron tomography (cryo-ET) imaging and live-cell fluorescence microscopy.

Architecture of the E. coli division site

Bacterial lamellae ~150–250 nm thick were generated by cryo-focused ion beam (cryo-FIB) milling 13 , 14 , 15 , 16 , 17 , 18 , 19 for in situ cryo-ET imaging (Extended Data Fig. 1 ). A total of 22 tilt-series of wild-type cells were acquired and three-dimensionally (3D) reconstructed (Fig. 1a , and Supplementary Tables 1 and 2 ). To gain better visualization of the sPG, nonlinear anisotropic diffusion (NAD) filtering was applied to denoise the cryo-electron tomograms (Fig. 1b and Supplementary Video 1 ). Densities corresponding to the OM, PG, and inner membrane (IM) were traced and 3D segmented (see Methods). Cells with an IM-IM distance >300 nm were classified as undergoing constriction. They had a V-shaped constriction with a relatively uniform invagination of the two membranes and an indented mesh of PG (Fig. 1a,b , Extended Data Fig. 2a and Supplementary Table 3 ). Cells classified as undergoing septation had an IM-IM distance <110 nm. They displayed a partial septum where the IM was more deeply invaginated than the OM, with an average difference of 138.5 ± 24.51 nm (Fig. 1a,b and Extended Data Fig. 2b ). Strikingly, the denoised tomograms showed an elongated, triangular wedge of PG close to the invaginating IM (Fig. 1b ). In cells at the final stages of cytokinesis, where IM fission was complete (Extended Data Fig. 2c ), two layers of PG signal comprising the septum were readily visible (Fig. 1b ). We performed subtomogram averaging to compare the envelope structure between the side wall and septum of dividing cells, which also showed two layers of PG signal within partial septa and a single layer of PG signal in the side wall (Extended Data Fig. 3a ).

a , Overview of different stages of cell division. Summed, projected central slices of cryo-electron tomograms visualizing different stages in division of wild-type E. coli are shown. Black arrowhead indicates the side of the division site displayed in ( b ). b , Top row: NAD-filtered cryo-electron tomograms visualizing the cell wall. Left panels show a 2D slice, right panels show the corresponding slice with segmentations for the observed PG signal in cyan, IM in green and OM in magenta (see Methods). White arrowheads indicate where the PG layer appears to thicken from one to two layers, and black arrowhead indicates the side of the division site shown in the schematic overview below. Bottom row: corresponding labelled summary diagrams. The left two bottom panels correspond to arrowhead-marked top division side rotated 90° to the left. Segmented PG signal is not indicative of specific glycan strand network. c – e , Representative time-lapse series from 3 biological replicates of wild-type E. coli expressing Pal-mCherry and ZipA-sfGFP as OM and IM markers, respectively, imaged at 30 °C on M9 supplemented with 0.2% casamino acids and d -glucose. Fluorescence signals were deconvolved (see Methods). The yellow triangle marks division sites used for line scans of fluorescence intensity (FI) profiles ( d ) and kymograph analysis of cytokinesis ( e ). f , Average constriction velocities of the IM and OM were derived from the slopes of the fluorescence signals in kymographs (see Methods). Black line indicates mean. Two-sided unpaired Mann-Whitney test; ** P  < 0.01; N  = 150 division kymographs. g , Instantaneous constriction velocities for ZipA (IM, green) and Pal (OM, magenta) are plotted against normalized cell width. Second order polynomial fits with 95% confidence intervals are shown. Scale bars: a and b , 100 nm; c , 2 µm; e , 200 nm (vertical) and 5 min (horizontal).

Source data

To investigate the mechanism of partial septum formation, we followed the constriction dynamics of each membrane. The IM was tracked using a superfolder green fluorescent protein (sfGFP) fusion to the IM-anchored Z-ring binding protein ZipA (ZipA-sfGFP), while constriction of the OM was followed using mCherry fused to the OM-lipoprotein Pal (Pal-mCherry) (Fig. 1c and Supplementary Video 2 ). The Pal-mCherry and ZipA-sfGFP signal distribution at the division site confirmed the deeper constriction of IM with respect to the OM (Fig. 1d ) observed by cryo-ET (Fig. 1a,b ). The invagination rate of each membrane was calculated from kymographs (Fig. 1e,f and Extended Data Fig. 4 ). We found that the IM constriction rate increased faster than linear as the septum closed, with an average rate of 64.26 ± 33.98 nm min −1 , in line with previous measurements 20 , 21 . The increase in the OM constriction rate during division was less pronounced than that of the IM (Fig. 1f,g and Extended Data Fig. 4b–d ). The different rates of change in constriction velocity between the two membranes account for the two membranes becoming increasingly separated as division proceeds, by 147 nm at late stages in cell division, which is in good agreement with our cryo-ET data. Thus, E. coli divides by a mixed constriction/septation mechanism, with the partial septum containing two layers of sPG signal (Fig. 1b and Extended Data Fig. 3a ).

sPG synthesis and remodelling defines septal architecture

We next determined how the architecture of the division site and the dynamics of its constriction are altered by mutations affecting sPG synthesis and remodelling. The essential PG synthase of the divisome is formed by FtsW and FtsI (FtsWI) 22 . Following Z-ring assembly, a regulatory pathway is initiated that activates sPG synthesis by this synthase 23 , 24 , 25 , 26 (Fig. 2a ). Activation is mediated in part via an interaction between FtsWI and the FtsQ-FtsL-FtsB (FtsQLB) complex 27 . Genetic evidence suggests that FtsQLB activation is stimulated by an essential peptide within FtsN 28 . Another domain of FtsN called SPOR concentrates the activation peptide at the division site through binding to sPG that was processed by PG amidases 28 , 29 , 30 . Amidases generate peptide-free (denuded) PG recognized by the SPOR domain as they split the sPG septum to promote OM constriction and daughter separation 31 . The interplay between sPG synthesis activation by FtsN and amidase processing bringing more FtsN to the division site promotes a positive feedback loop, the sPG loop, that has been proposed to drive cell division 28 . We imaged several mutants defective in this process (Supplementary Tables 4 and 5 ): (1) one lacking the SPOR domain of FtsN ( ftsN-∆SPOR ), (2) mutants defective for one ( ∆envC ) or both ( ∆envC ∆nlpD ) amidase activators 32 and (3) a mutant ( ftsL *) encoding a variant of FtsL that hyperactivates sPG synthesis 26 (Fig. 2a , Extended Data Figs. 3 b–f and 5 , and Supplementary Tables 1 – 3 and 6 ). All mutants displayed similar growth rates (Extended Data Fig. 6 ).

a , Schematic overview of the septal PG loop pathway for the activation of sPG synthesis (see text for details). b , Left: NAD-filtered cryo-electron tomograms of division sites in the indicated division mutants of E. coli shown as in Fig. 1b . Right: summary diagrams of the cell envelope architecture visualized. Black arrowheads indicate the side of the division site represented in the schemes. Segmented PG signal is not indicative of specific glycan strand network. c , Top: representative time-lapse series from 3 biological replicates of indicated E. coli division mutants expressing Pal-mCherry and ZipA-sfGFP as OM and IM markers, respectively, imaged as in Fig. 1 . Bottom: kymograph analysis and line scans of fluorescence intensity profiles of cytokinesis, from division sites marked with yellow triangles in the top row. d , Constriction velocities of the IM and OM were determined as in Fig. 1 . Black line indicates mean. Data from wild type are replotted from Fig. 1f for comparison. Brown-Forsythe and Welch ANOVA test with Dunnett’s correction for multiple comparisons, significance of differences is tested relative to wild type (wt); ** P  < 0.01, *** P  < 0.001, **** P  < 0.0001; NS, not significant ( P  = 0.09); N  = 150 (wt), 48 ( ftsN-∆SPOR ), 74 (∆envC ), 68 ( ftsL* ) kymographs. e , Instantaneous constriction velocities for IM (top) and OM (bottom) are plotted against normalized cell width. Second order polynomial fits with 95% confidence intervals are shown. See Extended Data Fig. 4b,c for individual instantaneous constriction velocity traces. Data from wild type are replotted from Fig. 1g for comparison. Scale bars: b , 100 nm; c , top row, 2 µm; bottom row kymographs, 200 nm (vertical), 5 min (horizontal).

Division sites from ftsN-∆SPOR cells resembled those observed previously for Caulobacter crescentus 33 (Fig. 2b ). By cryo-ET, there appeared to be greater coordination between IM and OM constriction throughout division (Fig. 2b , Extended Data Figs. 3 b–f and 5 , and Supplementary Video 3 ). Tracking of membrane constriction dynamics confirmed that the rate of constriction for the two membranes was nearly identical in the mutant (Fig. 2c–f , Extended Data Fig. 4b–e and Supplementary Video 2 ). As a result of the close opposition of IM and OM, the wedge of sPG observed in filtered tomograms was not as elongated as in wild-type cells, and separate plates of material forming ahead of the wedge were not observed (Fig. 2b ).

Cells defective for both amidase activators ( ∆envC ∆nlpD ) formed a near-complete septum in which the constriction of the IM was accomplished without much observable invagination of the OM (Fig. 2b , and Supplementary Figs. 3b –f and 4b–e ) . NAD-filtering revealed signal corresponding to sPG that was even more clearly discernible as two distinct plates of material than in wild-type cells (Figs. 1b and 2b ). Furthermore, the tomograms revealed a triangular wedge of PG material at the outer edges of the septa that was not previously observed in conventional EM analysis 34 and presumably serves as a roadblock to OM invagination (Fig. 2b and Supplementary Video 4 ). It was not possible to measure membrane constriction dynamics in live cells of the ∆envC ∆nlpD double mutant because it grew poorly when expressing the fluorescent markers. However, measurements in a mutant lacking only EnvC, the dominant amidase activator 32 , revealed a substantial disparity between IM and OM constriction rates (Fig. 2c–f and Supplementary Video 2 ). Cryo-ET data of the ∆envC strain showed a similar division site architecture as the double ∆envC ∆nlpD mutant (Extended Data Figs. 3 b–f and 5 ). Overall, the results indicate that impairing key components of the divisome converts the mixed constriction/septation division mechanism of E. coli to either a purely constriction or septation mode depending on the lesion.

Hyperactivated sPG biogenesis alters septal architecture

To determine the effects of hyperactivated sPG biogenesis on division site architecture, we imaged cells of the ftsL* mutant. Strikingly, cryo-ET revealed an altered architecture in which the signal corresponding to the wedge of sPG observed in wild-type and other mutant cells was missing, and the envelope at the leading edge of the invagination was 50.8% thinner than in wild-type cells (Fig. 2b and Extended Data Fig. 2d–f ). Conversely, the envelope in the nascent polar regions adjacent to the leading edge of the invagination was 12% thicker in ftsL* cells than in wild-type cells, with the bulged areas containing more PG than normal (Extended Data Fig. 2e and Supplementary Video 5 ). As expected from previous measurements 26 , the average constriction velocity of the IM was much greater than in wild-type cells (Fig. 2c–e , Extended Data Fig. 4b–f and Supplementary Video 2 ). Surprisingly, the average rate of OM constriction was much slower than that of the IM (Fig. 2c–f and Extended Data Fig. 4 ), a difference that would normally be expected to give rise to cells with partial septa. However, such architectures were not observed in the tomograms (Fig. 2b , and Extended Data Figs. 3 b–f and 5 ). The distances between the IM and OM remained relatively constant in all cells that were imaged (Fig. 2b , and Extended Data Figs. 2 a–c, 3 b–f and 5 ). This discrepancy is probably due to the PG binding activity of Pal causing the OM reporter to get stuck in the thicker PG that accumulates behind the closing septum and therefore to track poorly with the leading edge of the invaginating OM. Notably, upon closer inspection of kymographs of the ftsL* mutant, we noticed that one side of the cell constricted faster than the other (Fig. 2c ). When we compared the constriction velocity for each side of the division site and assessed the degree of anisotropy (Extended Data Fig. 4a ), ftsL* cells showed a higher but not statistically significant anisotropy score for both IM and OM constriction compared with other strains (Extended Data Fig. 7a–d ). Additionally, when cells were imaged in vertical orientation, the constriction of ftsL* cells was less isotropic than in wild-type cells, indicative of uneven closure of the division ring (Extended Data Fig. 7e ). Thus, circumvention of the normal controls regulating sPG biogenesis in the ftsL * mutant results in aberrant division site geometry and abnormal thickening of the envelope at the poles. These cells also lack an observable sPG wedge, which may destabilize the division site and help explain why these mutants were originally found to lyse at elevated temperatures 35 , 36 .

sPG degradation activates its synthesis

To better understand the mechanism(s) by which changes in division site architecture are caused by mutations altering divisome components, we measured the rates of sPG synthesis and degradation using two different cytological assays (Fig. 3a and Extended Data Fig. 8a ). The first assay used a pair of compatibly labelled fluorescent d -amino acids (FDAAs), YADA and HADA 37 , and the other used HADA and MurNAc-alkyne 20 . In both cases, cells were labelled extensively with an FDAA, pulsed with the second label for different lengths of time and then fixed before visualization. The intensity of the second label appearing at midcell after the pulse was used as a proxy for sPG synthesis. Additionally, the signal intensity of the first label before and after the pulse was used as a proxy for sPG degradation. Both assays yielded qualitatively similar results (Fig. 3a–g and Extended Data Fig. 8a–g ). The ftsL* mutant synthesized sPG faster than all other strains just as it had the fastest rate of IM invagination (Figs. 2d and 3b,c , and Extended Data Fig. 8b–d ). This result confirms that activated FtsQLB complexes indeed hyperactivate sPG synthesis, as suggested by recently reported effects on the dynamic motions of FtsWI 20 . Notably, the dual FDAA assay detected an increased amount of old PG at the division sites before the HADA pulse, and this material appeared to be relatively stable during the time course. Additionally, bright foci of old material were also observed at the poles of many cells after extended YADA labelling (Fig. 3b and Extended Data Fig. 8h,i ). This accumulation of old material probably corresponds to the thickened areas of cell wall in nascent poles observed by cryo-ET of the ftsL* mutant (Extended Data Figs. 5 and 8j,k ), reinforcing the conclusion that short-circuiting the normal controls governing sPG biogenesis not only leads to more rapid sPG synthesis and septal closure, but also aberrant accumulation of PG within the developing poles.

a , Labelling patterns observed for an FDAA pulse-chase experiment. New cell wall material is labelled with HADA (blue), while old material is stained with YADA (yellow). b , Representative images from 3 biological replicates of indicated strains after 2, 4 and 8 min pulses with HADA. Overlay images are provided in Extended Data Fig. 8l,m . c , d , Mean fluorescence intensity was measured at the division site for new ( c ) and old ( d ) PG. c , Data were fit to a linear regression to derive sPG synthesis rates. Data points represent median ± 95% confidence intervals. d , Reduction in old (YADA) fluorescence intensity was fit to a one-phase exponential decay curve. e , Mean septal PG hydrolysis rates were derived from decay curves in d . Points represent the average value of the three biological replicates and bars indicate mean + 1 s.d. N  = 1,054 (wt), 716 ( ftsN-∆SPOR ), 819 (∆envC ), 880 ( ftsL* ) cells. f , Side wall incorporation of new cell wall material (HADA fluorescence intensity) was measured after 8 min due to low signal intensities in earlier time points. Black line indicates median, one-way ANOVA with Dunnett’s correction for multiple comparisons, significance of difference is tested relative to wild type; NS, P  = 0.06; *** P  < 0.001, **** P  < 0.0001; N  = 103 (wt), 107 ( ftsN-∆SPOR ), 101 (∆envC ), 100 ( ftsL* ) cells. g , The ratio between sPG and side wall synthesis was calculated by dividing the mean HADA fluorescence intensity after the 8 min pulse. h – j , Labelling patterns observed for the pulse-chase experiment in cells with inhibited division by SulA expression ( h ). New cell wall material is labelled with Alexa488-labelled MurNAc-alkyne probes (yellow), while old material is stained with HADA (blue). Mean fluorescence intensity was measured along the side wall for both MurNAc-alkyne ( i ) and HADA ( j ) and fitted to a quadratic exponential Malthusian exponential growth function ( i ) or one-phase exponential decay ( j ). Data points represent median ± 95% confidence intervals. N  = 578 (wt), 456 ( ftsN-∆SPOR ), 427 (∆envC ), 501 ( ftsL* ) cells. k , Representative images from 3 biological replicates of indicated strains after 15, 20 and 30 min pulses with MurNAc-alkyne. Scale bar, 2 µm.

Both the ftsN-∆SPOR and ∆envC mutants displayed a reduced rate of sPG synthesis relative to wild-type cells (Fig. 3b,c and Extended Data Fig. 8c,d ). Although the sPG synthesis rates were similar, the two mutants differed in their rates of sPG degradation. The ftsN-∆SPOR mutant displayed relatively normal rates of sPG degradation, whereas ∆envC cells showed reduced turnover of sPG as expected for a mutant lacking an amidase activator (Fig. 3d,e and Extended Data Fig. 8e ). The combination of slower sPG synthesis with normal sPG degradation explains the well-coordinated constriction phenotype displayed by the ftsN-∆SPOR mutant in the cryo-ET analysis. The reduced rate of sPG synthesis in the ∆envC cells is notable because it indicates that proper sPG processing by the amidases is required for normal rates of sPG synthesis. This result along with the reduced rate of sPG synthesis observed for the ftsN-∆SPOR mutant provides strong support for the sPG loop model 28 .

sPG degradation is required for normal Z-ring formation

Cells lacking EnvC commonly displayed closely spaced sPG labelling consistent with the aberrant formation of adjacent division sites (Fig. 4a ). Accordingly, closer examination of the localization of Pal-mCherry in these cells revealed that double bands of the OM constriction marker occurred at an elevated frequency over wild-type or ftsN-∆SPOR cells (Fig. 4b,e,f ). These double constrictions were also observed in cryo-ET (Fig. 4d and Extended Data Fig. 5 ) and were typically placed within the cell body. We also observed constrictions near cell poles, generating what appeared to be minicells (Fig. 4c,g ). However, free minicells were not observed in the culture, suggesting that these aberrant poles were probably generated from a double constriction event within the cell body, one of which was aborted while the other completed division, generating a daughter with a polar constriction. Membrane blebs were also observed emanating from some of the developing septa of ∆envC cells, some of which appeared to lyse, suggesting there was a catastrophic failure in division.

a , Distribution of cell wall material in ∆envC cells was assessed by FDAA staining in 3 biological replicates. Images are sum-projections of a 1 µm spanning z -stack and were deconvolved. White arrowheads indicate double septa. b , Representative time-lapse series from 3 biological replicates of a ∆envC mutant expressing Pal-mCherry and ZipA-sfGFP as OM and IM markers, respectively. An example of double septum formation is shown. c , Examples of membrane blebbing (yellow arrowheads) and polar septa (blue arrowheads) formation are highlighted. d , Formation of double constrictions observed in cryo-electron tomograms of ∆envC ∆nlpD cells. Black arrowheads indicate constriction sites. e , The frequency of double septum formation was quantified from counting the number of Pal-mCherry doublets per cell. No Pal doublets were found in >10,000 cells for wild-type or ftsN-∆SPOR cells in 3 biological replicates (N.A., not applicable). Data are represented as median + 95% confidence interval. f , The distance between Pal doublets was measured manually using the line tool in Fiji. N  = 91 ( ∆envC ), 46 ( ftsL* ∆envC ) Pal doublets measured. g , The frequency of polar septa per cell was measured for the indicated strains. No polar septa were observed in >10,000 wild-type or ftsN-∆SPOR cells. Data are represented as median + 95% confidence interval. h – j , Three-dimensional maximum intensity renderings showing Z-ring condensation based on ZipA-sfGFP localization ( h ). The degree of Z-ring condensation was quantified from averaged fluorescence intensity projections from summed 3D volumes ( i ) or from 5 time points (corresponding to 10 min) of a time-lapse series ( j ) (see Methods). Insets: FWHM of the fluorescence signal, with data represented as boxplots; line represents median, error bars depict minimum–maximum range. Inserts show average fluorescence intensity projection at the septum. Significance was tested against wild type by one-way ANOVA with Dunnett’s correction for multiple comparisons: * P  < 0.05. N  = 100 (wt, ∆envC , ftsL* ∆envC , ftsN-∆SPOR ) Z-rings from 3 biological replicates. Averaged Z-rings are shown and colour-coded according to graphs. Scale bars: a – c , 2 µm; d , 200 nm; h , 2 µm; i and j , 200 nm.

The pattern of ZipA-sfGFP was also altered in the ∆envC mutant (Fig. 4h and Supplementary Video 6 ). ZipA binds FtsZ and is a Z-ring marker 38 . Many ZipA-sfGFP structures in ∆envC cells were diffuse and/or malformed, indicating a difficulty condensing into the tight Z-ring structure typical of normal cells (Fig. 4h and Extended Data Fig. 9 ). Consistent with this possibility, averaging the ZipA-sfGFP signals for a population of cells, either over a 10 min time window or over a 2 µm volume spanning midcell, showed that the fluorescence was more broadly distributed in the ∆envC mutant than in wild type or the other mutants (Fig. 4i,j ). Notably, this phenotype was not suppressed by combining the ftsL * mutation with ∆envC , indicating that it probably stems from the loss of sPG processing, not its collateral effect of reducing sPG synthesis (Fig. 4h–j ). A diffuse Z-ring phenotype has been observed for cells defective in FtsZ-binding proteins such as ZapA that are thought to bundle FtsZ polymers to condense the ring 39 , 40 . These results therefore suggest a previously unappreciated role for sPG hydrolysis by the amidases in Z-ring condensation and division site stability and/or placement.

Competition between elongation and sPG biogenesis

We took advantage of the PG labelling assays to quantify side wall PG synthesis (Fig. 3f and Extended Data Fig. 8f ) and found that it was inversely correlated with sPG synthesis. Side wall PG incorporation was highest in the ftsN-∆SPOR mutant, which had one of the lowest rates of sPG synthesis (Fig. 3c,f,g and Extended Data Fig. 8d,f,g ). Conversely, side wall PG synthesis was lowest in the ftsL * mutant that made sPG most rapidly (Fig. 3c,f,g and Extended Data Fig. 8d,f,g ). In support of a competition with cell division being responsible for the differing rates of side wall PG synthesis, the rates were found to be the same in all cells when cell division was blocked (Fig. 3h–k and Extended Data Fig. 8n–q ).

Another measure of cell elongation activity is the circumferential motion of the Rod complex associated cytoskeletal element MreB around the cell cylinder 41 , 42 , 43 . We tracked the motion of an mNeonGreen fusion to MreB in wild-type and mutant cells using a combination of structured Illumination microscopy and total internal reflection fluorescence (SIM-TIRF) imaging. Consistent with the sPG synthesis measurements, the total number of directionally moving MreB filaments per area was significantly reduced in ftsL* cells (Fig. 5a,b , Extended Data Fig. 10a,b and Supplementary Video 7 ), which had an increased cell width (Extended Data Fig. 10c,d ) indicative of reduced Rod complex activity 44 . All mutants displayed a similar density of directionally moving MreB filaments following the inhibition of cell division (Extended Data Fig. 10e,f and Supplementary Video 7 ), providing further support for a competition between the processes of elongation and division.

a , MreB dynamics were followed by SIM-TIRF in indicated strains (see Methods). Time-lapse series were sum projected and overlayed with single-particle tracking results from TrackMate and 3D-SIM Pal-mCherry reference images. The Pal-mCherry signal serves to identify constricting cells. Early division site (yellow arrowheads) displayed Pal foci that were resolvable as two distinct foci, whereas late division sites (blue arrowheads) displayed a continuous Pal signal across the cell, indicative of complete or near-complete cytokinesis. b , Directionally moving MreB tracks were filtered by MSD analysis (see Methods), represented as boxplots (line indicates median; error bars depict minimum–maximum range) and normalized by cell area. Significance in each group was tested against wild type by one-way ANOVA with Dunnett’s correction for multiple comparisons: * P  < 0.05, ** P  < 0.01; NS, P  ≥ 0.05. N  = 30 (wt, ftsN-∆SPOR, ∆envC, ftsL* ) time-lapse series from 3 biological replicates. c , Representative phase-contrast micrographs showing segmented cells in ‘Morphometrics’ for the indicated division mutants. d , Summed, projected central 3D slices through cryo-electron tomograms of indicated strains visualizing cell poles. Black arrowheads indicate 3D-rendered pole. The corresponding 3D-volume renderings show polar curvature determined by shape index (see Methods). e – g , Polar curvature was measured by the two highest points of positive cell outline curvature ( f ), while constriction curvature was assessed by measuring the opposing contour-matched lowest curvature values at the division site ( g ) using Morphometrics and normalized to cell width (see Methods). Polar and division site curvatures are negatively correlated ( R 2  = 0.27) ( e ). Data are represented as mean ± s.d. For f and g , significance was tested against wild type by one-way ANOVA with Dunnett’s correction for multiple comparisons: *** P  < 0.001, **** P  < 0.0001; NS, P  = 0.057. N  = 460 (wt), 999 ( ftsL* ), 292 ( ftsN-∆SPOR ), 164 (∆envC ) cells from 3 biological replicates. Scale bars: a , 1 µm; c , 2 µm; d , summed projection images, 200 nm and 3D renderings, 100 nm.

Notably, the interplay between cell elongation and division impacted the geometry of the division site and the shape of the daughter cell poles (Fig. 5c,d ). The ftsN-∆SPOR mutant, which elongates more rapidly and constricts slower, displayed an elongated division site and a shallower OM invagination angle at midcell as compared with wild-type cells (Fig. 5e–g ). This altered constriction geometry was also observable by cryo-ET and correspondingly gave rise to daughter cells with pointier poles than wild-type cells (Fig. 5c–g ). On the other hand, the rapidly constricting ftsL* mutant formed daughter cells with relatively blunt cell poles (Fig. 5c–g ).

We reasoned that the variation in division site and polar geometry among the different strains could be related to the activity of the Rod complex at or near the division site. The number of directionally moving MreB filaments in proximity (≤200 nm) to cell constrictions was therefore quantified (Fig. 5a,b and Supplementary Video 8 ). Such filaments were readily observed to pass through division sites in both early and late pre-divisional cells in all strains tested. Notably, however, the ftsN-∆SPOR mutant displayed more MreB tracks at the division site at late stages of division than all other strains, and the ftsL* mutant showed the least number of total MreB tracks at the division site (Fig. 5b ). Thus, the density of MreB tracks at the division site for these cells correlates well with the steepness of the constriction site and the extent of cell pole elongation observed for the different strains. The outlier was the ∆envC mutant, which had an inverted trend of having fewer directionally moving MreB tracks at early division stages than at later points (Fig. 5b ). We suspect that this change is due to the defect in sPG splitting, which causes a steep curvature of the inner membrane at early points in division that is probably unfavourable for MreB localization 45 , 46 . However, at later stages when sPG processing eventually allows for slow constriction of the OM, this curvature probably becomes more favourable for MreB localization, allowing elongation to occur near the division site to generate a shallow constriction such as that of the ftsN-∆SPOR mutant. Overall, these results not only provide strong support for a competition between the PG biosynthetic machineries involved in cell elongation and division, but also highlight the potential for this competition to define the morphology of the daughter cell poles.

Architecture of the sPG layer

Here we combined cryo-FIB milling with cryo-ET to visualize the division site of E. coli in situ. In cells just starting to constrict, all three envelope layers appeared to be invaginating in concert, and little change in the sPG relative to the side wall PG was evident. However, the speed of IM invagination and sPG synthesis increases faster than PG splitting and OM constriction, leading to the formation of a partial septum (Fig. 6a ) similar to that previously observed in fixed samples 9 , 10 . In NAD-filtered tomograms, a triangular wedge of what is likely to be sPG is observed at the lagging edge of the septum closest to the tip of the invaginating OM (Fig. 6a ). The wedge thins as it approaches the leading edge of the closing IM. In this narrow portion of the septum, two dense tracks of material are often discernible, which correspond to the PG layers that will eventually fortify the daughter cell poles. In ftsN-∆SPOR cells with reduced sPG synthesis activity and slower IM constriction, a more uniform constriction of all envelope layers is observed, generating a division site architecture that resembles that of C. crescentus 33 (Fig. 6b ). However, in cells defective in sPG splitting, OM constriction is almost completely blocked and a Gram-positive-like septum is formed, with two visible tracks of PG reminiscent of the two tracks observed in the developing septa of Staphylococcus aureus 7 (Fig. 6b ). These results suggest that the activity of the same basic cell division machinery can generate different septal architectures observed in diverse bacteria. All that may be required is to change the relative activities of the sPG synthesis and remodelling systems.

a , Wild-type E. coli divides via a mixed constriction-septation mechanism in which a partial septum with two discernible plates of sPG is formed at later stages of the division process. A wedge structure is observable at the lagging edge of the septum where the dual layers of PG signal of the developing septum meet the single-layered signal of the side wall. Although not clearly resolved in the tomograms, we assume that the two layers of PG signal within the septum are probably connected by additional PG material (drawn as crosshatches). b , A constrictive mode of cell division is observed for the ftsN-∆SPOR mutant, where OM and IM invaginate at similar velocities due to lower sPG synthesis rates. The result is a V-shaped constriction that is similar to that formed by the distantly related Gram-negative bacterium C. crescentus . In contrast, inhibition of sPG hydrolysis causes a temporal separation of IM and OM constriction, leading to septation. These septa as well as the partial septa in wild-type cells are reminiscent of the Gram-positive bacterium S. aureus , which also displays two distinctive plates of sPG within its septa. c , The activities of the two major synthetic cell wall machineries, the Rod complex and the divisome, are anti-correlated probably due to competition for limited substrate (lipid II). The balance of their relative activities determines the shape of the cell division site and the resulting poles they form. Cells with higher Rod complex activity are thinner and form pointier poles, while cells with elevated divisome activity are shorter and wider, with blunt poles.

In cells defective in sPG processing by the amidases, the sPG wedge structure is more prominent than in wild-type cells and it appears to impede the invagination of the OM. We thus infer that amidases process this structure to allow constriction of the OM (Fig. 6a ). Furthermore, because the sPG wedge is observed in deeply constricted wild-type cells as well as unconstricted amidase activation mutants, we suspect that the structure is dynamic, with its lagging edge being degraded as new wedge material is deposited at the leading edge. Such a spatial separation of synthesis and degradation would allow the sPG wedge to move in a treadmill-like fashion ahead of the OM as the septum closes.

The enzymes responsible for creating the sPG wedge remain to be identified, but our results with the ftsL * mutant suggest that it is not made by FtsWI. This mutant is thought to hyperactivate FtsWI 20 , 24 , 25 , 26 , 27 . Therefore, if the wedge were produced by the FtsWI synthase, the ftsL * mutant would be expected to produce a thicker or otherwise larger wedge. Instead, it lacks a wedge altogether, suggesting that enhanced FtsWI activity disrupts biogenesis of the sPG wedge by other synthases. An attractive candidate for this additional synthase is the class A penicillin-binding protein (aPBP) PBP1b. Inactivation of PBP1b is synthetically lethal with defects in FtsWI activation. The affected mutants were found to lyse due to septal lesions, suggesting that this aPBP promotes division site stability 24 , 47 . The location of the wedge at the lagging edge of the division site closest to the OM is also consistent with a role for PBP1b in its construction, given that this enzyme requires an OM lipoprotein for activity 48 , 49 . Thus, the outer fork of the division site where the wedge is located is the only place where aPBPs would be predicted to be functional. Although further work will be required to test this model, it provides an attractive explanation for the division of labour between the aPBP and FtsWI synthases during constriction, with the FtsWI synthase promoting ingrowth of the PG layer and the aPBPs providing backfill to stabilize the septum and prevent lysis.

The sPG activation loop

Our results support the proposal that FtsN and the amidases cooperate in a positive feedback loop that promotes sPG synthesis 28 . In addition to stimulating sPG synthesis, our results indicate that the sPG activation pathway also appears to be important for normal septal architecture. The ftsL * mutant hyperactivates the FtsWI synthase and eliminates the strict FtsN requirement for sPG biogenesis 26 . This short-circuiting of the normal division activation pathway not only causes the loss of the sPG wedge structure, but also promotes the aberrant accumulation of PG within the developing poles. Whether this accumulation results from inappropriate activation of PG synthesis by FtsWI or PBP1b, the improper turnover of the deposited material, or some combination of the two remains unknown. Nevertheless, what is clear is that bypassing the normal controls involved in sPG activation has adverse consequences on the architecture of the poles that are formed. We therefore infer that the normal divisome activation pathway serves an important function in coordinating different activities of the machinery to ensure that division is successfully completed once it is initiated and that the polar end products have a uniform surface.

PG hydrolysis and the Z-ring

Our results have uncovered an unexpected connection between the activation of sPG processing by the amidases and the Z-ring structure, suggesting that there is feedback to the Z-ring from events downstream of sPG synthesis activation (Fig. 6a ). Z-rings were found to be poorly condensed in mutant cells lacking the amidase activator EnvC (Fig. 4h–j ) . Additionally, closely spaced constrictions or areas of sPG biogenesis were also observed at an elevated frequency in these cells, suggesting that division sites are unstable and fail before they complete the division process (Fig. 4a–g ). Taken together, these results suggest the counterintuitive notion that sPG degradation by the amidases is required to stabilize the divisome, most probably via a positive influence on Z-ring condensation. Given that the amidases act on sPG in the periplasm, they are unlikely to directly modulate FtsZ activity. Rather, their effect is probably mediated through SPOR domain proteins such as FtsN and DedD that bind the amidase-processed glycans 29 , 30 . These proteins have transmembrane domains and N-terminal cytoplasmic tails, which in the case of FtsN is known to associate with the FtsZ-binding protein FtsA 50 , 51 . Thus, the status of sPG biogenesis in the periplasm could be communicated to the Z-ring in the cytoplasm using the binding of SPOR domain proteins to denuded glycans as a proxy. Whether the effect might be mediated simply by concentrating the cytoplasmic domains of SPOR proteins at the division site to modulate the activity of FtsZ-binding proteins or via more complex mechanisms requires further investigation, but the emerging picture is that the divisome activation pathway is not a one-way street from Z-ring formation to sPG synthesis and processing. The Z-ring probably also receives return stabilizing/activating signals from the PG biogenesis machinery.

Cell shape and the balance between division and elongation

The idea that the cell elongation and division machineries may be in competition with one another has been discussed in the field for some time 52 , 53 . However, it has only been recently that evidence for such a completion has been presented 47 , 52 , 53 . Here we used several independent assays to demonstrate that septal and side wall PG synthesis rates are inversely correlated to each other, providing strong support for antagonism between the activities of the elongation and division systems, which most probably stems from a competition for the limited supply of the lipid II PG precursor. Importantly, our results indicate that this competition does not just affect cell width or how long or short cells are. It also influences the geometry of the septum and the shape of the daughter cell poles. Thus, modulation of the relative activities of the elongation and division systems is likely to play an important role in generating the diversity of shapes observed among different bacteria.

Media, bacterial strains and mutagenesis

Indicated strain derivatives of E. coli MG1655 used in this study are listed in Supplementary Tables 4 and 5 . Bacteria were grown in LB (1% Tryptone, 0.5% yeast extract, 0.5% NaCl) or M9 media 54 each supplemented with 0.2% d -glucose and casamino acids. For selection, antibiotics were used at 10 µg ml −1 (tetracycline), 25 µg ml −1 (chloramphenicol) and 50 µg ml −1 (kanamycin, ampicillin). Mutant alleles were moved between strains using phage P1 transduction. If necessary, the antibiotic cassette was removed using FLP recombinase expressed from pCP20 55 . All mutagenesis procedures were confirmed by PCR.

Cryo-EM specimen preparation

Extended Data Fig. 1 summarizes the cryo-FIB/cryo-ET pipeline utilized in this study. Bacterial strains were grown overnight in LB media, back diluted 1:1,000 and incubated with shaking at 37 °C and 250 r.p.m. to optical density (OD) 600  = 0.3. Cells were collected by centrifugation (2 min, 5,000 ×  g , r.t.) and resuspended in LB media to a final OD 600  = 0.6. This cell suspension (3 µl) was applied to Cflat-2/1 200 mesh copper or gold grids (Electron Microscopy Sciences) that were glow discharged for 30 s at 15 mA. Grids were plunge-frozen in liquid ethane 56 with an FEI Vitrobot Mark IV (Thermo Fisher Scientific) at r.t., 100% humidity with a waiting time of 10 s, one-side blotting time of 13 s and blotting force of 10. Customized parafilm sheets were used for one-side blotting. All subsequent grid handling and transfers were performed in liquid nitrogen. Grids were clipped onto cryo-FIB autogrids (Thermo Fisher Scientific).

Cryo-FIB milling

Grids were loaded in an Aquilos 2 Cryo-FIB (Thermo Fisher Scientific). The specimen was sputter coated inside the cryo-FIB chamber with inorganic platinum, and an integrated gas injection system was used to deposit an organometallic platinum layer to protect the specimen surface and avoid uneven thinning of cells. Cryo-FIB milling was performed on the specimen using two rectangular patterns to mill top and bottom parts of cells, and two extra rectangular patterns were used to create micro-expansion joints to improve lamellae instability 57 . Cryo-FIB milling was performed at a nominal tilt angle of 14°−18°, which translates into a milling angle of 7°−11° 58 . Cryo-FIB milling was performed in several steps of decreasing ion beam currents ranging from 0.5 nA to 10 pA and decreasing thickness to obtain 100–200 nm lamellae.

All imaging was done on an FEI Titan Krios (Thermo Fisher Scientific) transmission electron microscope operated at 300 KeV and equipped with a Gatan BioQuantum K3 energy filter (20 eV zero-loss filtering) and a Gatan K3 direct electron detector. Before data acquisition, a full K3 gain reference was acquired, and ZLP and BioQuantum energy filters were finely tuned. The nominal magnification for data collection was ×42,000 or ×33,000, giving a calibrated 4 K pixel size of 2.193 Å and 2.565/2.758 Å, respectively. Data collection was performed in the nanoprobe mode using the SerialEM 59 or Thermo Scientific Tomography 5.3 software. The tilt range varied depending on the lamella, but was generally from −70° to 70° in 2° steps following the dose-symmetric tilt scheme 60 . Tilt images were acquired as 8 K × 11 K super-resolution movies of 4–8 frames with a set dose rate of 1.5–3 e −  Å −1  s −1 . Tilt series were collected at a range of nominal defoci between −3.5 and −5.0 µm and a target total dose of 80–180 e −  Å −2 (Supplementary Table 1 ).

Cryo-ET image processing

Acquired tilted super-resolution movies were motion corrected and Fourier cropped to 4 K × 5 K stacks, using ‘framealign’ from IMOD 61 . Tilt series were aligned using ‘etomo’ in IMOD 62 and ‘Dynamo’. Contrast transfer function (CTF) estimation was performed in IMOD. CTF correction was performed using the ‘ctfphaseflip’ programme in IMOD 63 . CTF-corrected unbinned tomograms were reconstructed by weighted back projection with and without a SIRT-like filter and subsequently 2x, 4x and 8x binned in IMOD 62 .

Bandpass filtering and summed projection of cryo-tomogram slices were performed in Dynamo 64 , 65 , 66 , 67 complemented with customized MATLAB scripts. Gaussian and NAD-filtering were performed in Amira (Thermo Fisher Scientific) for visualization purposes. NAD-filtering was applied using the command ‘Anisotropic Diffusion’ in 3D mode for 5 iterations. Gaussian filtering was done by applying the command ‘Gaussian Filter’ under 3D mode with a kernel size factor of 3. Whole 3D-volume FFT filtering was performed in IMOD.

Segmentation

Segmentation was performed on FFT filtered and NAD-filtered tomograms using Amira (Thermo Fisher Scientific) by non-biased semi-automatic approaches. Manual annotation was required every 10 slices, then Amira’s interpolation function was applied to automatically trace slices in between. Annotation was done in two-dimensional (2D) slices where features of interest were visible by eye. The segmented PG signal is not indicative of specific glycan strand network, but rather serves as a visual guide to relevant cell wall features.

Three-dimensional pole curvature rendering was performed in Amira by applying the command ‘Curvature’ on the basis of the triangulated 3D mesh and ‘Shape Index’ as implemented in Amira 68 . Shape index (SI) computes the surface scalar field, which is calculated as

where C 1 and C 2 are the two principal curvatures. Shape index ranges from −1 to 1, negative values indicate negative curvature, positive values indicate positive curvature and values close to 0 indicate flatness of the surface. Values are normalized with respect to neighbouring triangles’ SI values 68 (Fig. 5d ).

Quantification of cryo-ET data

Division site dimensions.

Summed projection images of cryo-ET tomograms were used to quantitatively measure cell dimensions at the division site 69 . Measurements were performed in Fiji 70 using the ‘point to point’ measuring tool. Measurements were from IM to IM and from OM to OM.

Periplasmic space

Measurements of periplasmic space thickness were performed from the centre of the OM to the centre of the IM in the cell areas referred to here as ‘side wall’, ‘pole’ and ‘curve’ as well as the invagination tip of the OM to the IM at the constriction division stage. Measurements from centre to centre of opposing IMs were performed in the cell area defined in this study as the septum (Supplementary Figs. 2 and 5 ) . We used a customized macro in Fiji that measures 30 Euclidean distances from surface-to-surface areas 71 in nm, for example, from IM to IM at the septum and from IM to OM at the rest of the areas (side wall, pole, curve and initiation). For these 30 single measurements, the mean was calculated, yielding a final single value per defined subcellular localization, for example, septum, curve, pole and side wall.

Subtomogram averaging

Subtomogram averaging was performed in Dynamo 64 . From the full wild-type cryo-ET data set, particles were identified using ‘dtmslice’ interface in Dynamo 66 , 67 , 72 . In 4x-binned tomograms, subtomograms with a size of (777.6) 3  Å were extracted from 4x-binned tomograms. Initial angles were assigned following the normal of the IM. A starting reference generated from a random set of particles was used for both side wall and septum particles. A total of 16 iterations were used to align particles and obtain final averages. Final averages were generated from 8,076 subtomograms for the side wall and 212 particles for the septum. Notice that side wall regions were much more abundant in the cell than septum regions. EM densities were visualized in Chimera 73 .

Sample preparation for live cell imaging

Overnight cultures of indicated E. coli strains were grown in LB supplemented with appropriate antibiotics at 37 °C. The next day, cells were collected by centrifugation (2 min, 5,000 ×  g , r.t.) and washed 2× with M9 medium. Day cultures were back diluted (1:1,000) and grown in M9 (0.2% d -glucose, 0.2% casamino acids) supplemented with 50 µM Isopropyl β-D-1-thiogalactopyranoside (IPTG) and appropriate antibiotics at 30 °C until OD 600  = 0.2–0.4. For filamentation experiments, SulA was produced from pNP146 74 by the addition of 0.2% l -arabinose during the last 10 min of the incubation period. Cells were collected (2 min, 5,000 ×  g , r.t.) and resuspended in 1/10th of the original volume. Two microlitres of this cell suspension were added onto a 1% (w/v) agarose in M9 (0.2% d -glucose, casamino acids) pad supplemented with 50 µM IPTG and covered with a #1.5 coverslip. For filamentation experiments, the agar pad was also supplemented with 0.2% l -arabinose.

Live-cell imaging

All samples were imaged on a Nikon Ti-E inverted widefield microscope equipped with a fully motorized stage and perfect focus system. Images were acquired using a 1.45 NA Plan Apo ×100 Ph3 DM objective lens with Cargille Type 37 immersion oil. Fluorescence was excited using a Lumencore SpectraX LED light engine and filtered using ET-GFP (Chroma, 49002) and ET-mCherry (Chroma, 49008) filter sets. Images were recorded on an Andor Zyla 4.2 Plus sCMOS camera (65 nm pixel size) using Nikon Elements (v5.10) acquisition software. For subsequent deconvolution procedures, three 200 nm spaced Z -planes were acquired for both fluorescence channels using 100% LED output power and 50 ms exposure. Temperature was maintained at 30 °C using a custom-made environmental enclosure. After a 20 min acclimatization period, cells were imaged at a 2.5 min acquisition frame rate for a total observation time of 1–4 h.

Image processing for fluorescence microscopy

First, time-lapse series and Z -stacks were drift corrected using a customized StackReg plugin in Fiji 70 , 75 . Subsequently, fluorescence images were deconvolved using the classical maximum likelihood estimation algorithm in Huygens Essential v19.10 (SVI), employing an experimentally derived point spread function (PSF) from 100 nm TetraSpeck beads (Thermo Fisher Scientific). Image reconstruction was performed over 50 iterations with a quality threshold of 0.01 and a signal-to-noise ratio set to 20 for live-cell imaging and 40 for fluorescent cell wall probes in fixed samples. Background removal was set to 0 to preserve fluorescence intensity values best among different images. Chromatic aberrations between different fluorescent wavelengths were post-corrected using the chromatic aberration corrector in Huygens from the TetraSpeck bead template. The same image reconstruction parameters and chromatic aberration templates were applied to images that were compared to each other. Last, reconstructed fluorescence images were merged back to phase-contrast images and rendered for figure or movie display with Fiji.

Measuring cell envelope constriction dynamics

Fluorescent fusions to IM-anchored protein ZipA and OM-lipoprotein Pal allowed us to determine the respective positions of the different cell envelope layers during division. These cell envelope fiducial markers accumulate specifically during cytokinesis at the division site, which was critical for the generation of kymographs. Constriction dynamics of IM and OM were derived from kymographs generated using the Fiji plugin ‘KymographClear’ 76 and automatically split into forward and reverse trajectories using Fourier filtering. This filtering step allowed us to measure the constriction rate for each side independently. Constriction kinetics were derived by automatically extracting the fluorescent trajectories for ZipA and Pal using ‘KymographDirect’ 76 (Extended Data Fig. 4a ). Anisotropy of the division process was determined by taking the ratio of the constriction velocities between the forward and reverse trajectories. Only cells where the division site displayed minimal signs of displacement except for constriction were analysed to eliminate confounding effects on the analysis by excessive cell movement (for example, pushing). This manual exclusion resulted in the rejection of approximately 15–20% of the cell division events. Applying these procedures, we found the constriction rate of the OM to be increasing over time, in contrast to a previously reported constant rate 77 . This might be explained by different image analysis procedures (for example, kymographs vs width measurements, Pal-mCh marker vs a combination of phase-contrast and FM4-64 dyes).

Measuring division site circularity of vertically imaged cells

For vertical imaging of bacterial cells undergoing division, similar procedures as described previously 78 , 79 were applied. A silicon wafer containing 5.5 µm long and 1.5 µm wide photo-resist pillars was generated following high aspect ratio photolithography procedures with an adhesion layer. The dimension of these pillars reaches the practically feasible aspect ratio for photolithography designs and thus impedes increasing pillar length without concomitantly increasing width, precluding use of elongated or chaining division mutants for this imaging mode. A modified silanization surface treatment with plasma cleaning was applied to increase the surface hydrophobicity of the silicon wafer to minimize agarose accumulation. Agarose micro holes were generated by pouring degassed 6% agarose (w/v) in H 2 O on the silicon wafer. Agarose was allowed to solidify for 40 min at r.t., was peeled off, cut into 5 ×5 mm pieces, and incubated in M9 medium supplemented with 0.2% d -glucose, casamino acids, 25 µg ml −1 chloramphenicol and 50 µg ml −1 ampicillin overnight.

Cells were grown as described for sample preparation for live-cell imaging and added on agarose pads. Cells that were not trapped in micro holes were washed off gently using 1 ml of growth medium. Five micrometre spanning Z -stacks (at a 200 nm step size) were acquired and subsequently deconvolved.

Circularity quantification was carried using the software package ‘Morphometrics’ 80 . Fluorescence signals were segmented using Laplacian algorithm in combination with the peripheral fluorescence setting. Circularity ( C ) is calculated in Morphometrics as:

where P is the perimeter and A is the area enclosed by the circle and is a dimensionless measure. A perfect circle displays a circularity of 1, while increasing values correspond to less circular objects. Cells that were trapped tilted in agar holes were manually excluded from the analysis (15 out of 573 analysed cells).

Measuring Z-ring condensation from time-lapse data

Condensation of cytoskeletal elements was addressed using previously described procedures 40 . Briefly, five frames (corresponding to 10 min) from recorded time-lapse series were sum-projected in Fiji. Z-rings in these sum-projected images were then aligned along the length axis and average-intensity-projected into a single image. Fluorescence intensity was measured across the full width along the horizontal axis of the averaged projection image. Intensity values were normalized and their corresponding full width at half maximum (FWHM) values were calculated in MATLAB.

Measuring Z-ring condensation from 3D data

Similar procedures as outlined for measuring Z-ring condensation in time-lapse series were applied. Two micrometre spanning Z -stacks (at a 200 nm step size) were acquired to capture a full 3D view of a cell. Images were restored in Huygens as described above. Image volumes were sum-projected into a single plane, Z-rings extracted, aligned and averaged as described above. Fluorescence intensity profiles were measured identically as for time-lapse data. Snapshots for 3D maximum intensity projections were rendered in Huygens.

Measuring cell wall synthesis rates by biorthogonal MurNAc-alkyne probes

Septal cell wall synthesis rates were measured as described previously 81 , 82 . MurNAc-alkyne was purchased as a custom synthesis product from Tocris following the procedures of ref. 81 . All experiments were carried out in ∆murQ background and in the presence of pCF436 83 for IPTG-inducible expression of AmgK and MurU. Filamentation was induced by expressing the FtsZ antagonist sulA from arabinose-inducible plasmid pNP146 74 . Overnight cultures were back diluted 1:1,000 into fresh LB containing 15 µg ml −1 gentamycin. Cells were grown at 37 °C until OD 600  = 0.4. Subsequently, 1.5 ml of cells were collected (2 min, 5,000 ×  g , r.t.) and resuspended in 300 µl LB containing 1 mM IPTG and 0.5 mM HADA to label all cell wall material with FDAAs. For filamentation experiments, SulA expression was induced by the addition of 0.2% l -arabinose. Samples were incubated by rotating at 37 °C for 30 min. Endogenous UDP-MurNAc production was inhibited by the addition of 200 µg ml −1 fosfomycin. After 10 min incubation, cells were washed twice in 1.5 ml LB, 1 mM IPTG and 200 µg ml −1 fosfomycin. Next, cells were incubated for 15 min in the presence of 0.2% (w/v) MurNAc-alkyne, 1 mM IPTG and 200 µg ml −1 fosfomycin at 37 °C. Cells were fixed using ice-cold 70% (w/v) ethanol for 20 min at 4 °C. Next, cell pellets were washed 3× with 1x PBS. Biorthogonal MurNAc-alkyne probes were labelled by click chemistry using 5 µM Alexa488 azide substrate according to the manufacturer's instruction. Samples were stored in 20 µl PBS at 4 °C and imaged within 48 h of the labelling experiment.

Samples were imaged on a Nikon Ti2-E inverted widefield microscope equipped with a Lumencor Spectra III light engine, Semrock dichroics (LED-CFP/YFP/mCherry-3X-A-000, LED-DA/FI/TR/Cy5/Cy7-5X-A-000) and emission filters (FF01-432/36, FF01-515/30, FF01-544/24). Images were recorded using a 1.45 NA Plan Apo ×100 PH3 oil objective with Olympus Type F immersion oil and a pco.edge 4.2bi back illuminated cooled sCMOS camera using Nikon Elements 5.2.

One micrometre spanning Z -stacks (separated by 200 nm) were acquired and subsequently deconvolved as described above. Z -stacks were sum-projected using Fiji. De novo septal PG synthesis was assessed by measuring the mean fluorescence intensity of NAM-Alexa488 along the division site using the line tool (width, 3 pixels). Levels of cell wall hydrolysis were assessed by measuring the overall reduction in HADA fluorescence as compared to baseline signal intensity derived from fixing cells before MurNAc-alkyne chase. Reduction in fluorescence intensity of FDAAs is indicative of cell wall remodelling mediated by amidases, endopeptidases or transglycosylases.

Measuring cell wall remodelling by FDAA incorporation

For FDAA pulse-chase experiments, cells grown overnight were back diluted 1:1,000 in fresh LB and grown until OD 600  = 0.4 at 37 °C. For the filamentation experiment, sulA was expressed from pNP146 74 by the addition of 0.2% l -arabinose to cultures during the last 10 min of the incubation period. Subsequently, 1.5 ml of cells were collected (2 min, 5,000 ×  g , r.t.) and resuspended in 300 µl LB containing 0.5 mM YADA. Samples were incubated while rotating at 37 °C for 40 min. Cells were washed once in 1.5 ml LB and resuspended in 300 µl LB containing 0.5 mM HADA. Samples were incubated at 37 °C for either 2 min, 4 min or 8 min before immediate fixation with 70% ethanol. After fixation, cells were washed 3× in PBS, stored in the dark at 4 °C and imaged within 48 h. The same image acquisition and analyses procedures were carried out as described for MurNAc-alkyne probes. Fluorescence intensity values were fit to a linear regression for HADA and an exponential one-phase decay for YADA. Levels of cell wall hydrolysis were assessed by subtracting the average fluorescence intensity from cells fixed before chase (0 min) and the respective time point, and fit to a linear regression model. Reduction in fluorescence intensity of FDAAs is indicative of cell wall remodelling mediated by amidases, endopeptidases or transglycosylases. In addition to the division site, fluorescence intensity measurements were also performed along the side wall and polar region of the cells at the 8 min time point. For filamenting cells, HADA fluorescence intensity values were fit to a Malthusian exponential equation, assuming cells keep elongation at the same rate before SulA induction.

Bulk growth curve measurements

Overnight cultures of indicated E. coli strains were grown in LB supplemented with appropriate antibiotics at 37 °C. The next day, cells were collected by centrifugation (2 min, 5,000 ×  g , r.t.) and washed 2× with the respective growth medium (M9 or LB). Day cultures were back diluted (1:1,000) and grown in the respective media supplemented with corresponding IPTG concentration (50 µM for ZipA-sfGFP induction, 1 mM for AmgK/MurU expression) and appropriate antibiotics at 30 °C until OD 600  = 0.3. Cells were collected (2 min, 5,000 ×  g , r.t.) and resuspended to an initial OD 600 of 0.01 in a final volume of 100 µl. Growth curves were measured in a Tecan M-plex 96-well plate reader by OD 600 read out. Plates were incubated with shaking at 30 °C for a total of 18 h.

Cell shape quantification analyses

Bacterial cells were segmented and analysed from phase-contrast images using the software package ‘Morphometrics’ 80 . Results from Morphometrics were post-processed using customized MATLAB scripts to exclude erroneously segmented cell debris in live-image data on the basis of area. Cell width, length and pole curvature per segmented cell were directly extracted from Morphometrics. Since curvature ( \(k = \frac{1}{r}\) , where r is the radius of the cell cylinder) is dependent on the cell cylinder width, curvature values were normalized by multiplying half-cell width to each respective curvature value. Thus, spherical poles display curvature values of k  = 1, while pointy (elongated) poles display elevated curvature values ( k  > 1) and flat (shortened) poles display reduced curvature values k  < 1, respectively. We obtained division site curvature from both sides of the cell at the invagination site. The invagination site is defined as the narrowest segment of the cell, for example, lowest cell width value, that presents negative curvature on both sides of the cell body. Division site curvature was normalized to the half-cell width of the invagination site.

SIM-TIRF microscopy and MreB tracking

Samples were prepared as described for live-cell imaging. To block cell division, sulA was expressed from pNP146 74 by the addition of 0.2% l -arabinose during the last 10 min of the incubation period. Cells were added to high precision #1.5 coverslips (Marienfeld) and placed on a 1% (w/v) agarose pad in M9 (0.2% d -glucose, casamino acids, supplemented with 0.2% l -arabinose for filamentation experiments) and imaged at room temperature on a Nikon Ti2 N-SIM microscope equipped with N-SIM spatial light modulator illuminator, TIRF Lun-F laser combiner with 488 and 561 nm laser lines, an N-SIM 488/561 dual band dichroic mirror, SR HP Apo TIRF ×100 1.5 NA oil objective with automated correction collar and a Hamamatsu Orca Flash 4.0 camera attached to a Cairn Research Twimcam splitter with an ET525/50m or an ET605/70m emission filter (for MreB-sw-mNeonGreen or Pal-mCherry fusion, respectively). The refractive index of the immersion oil (1.512) (GE Healthcare) was optimized for MreB-sw-mNeonGreen signal and corrected using the automated correction collar for the Pal-mCherry fusion. Alignment of the 488 and 561 lasers for SIM-TIRF and 3D-SIM, and of the N-SIM optics and illumination was performed before each experiment at the image plane. First, a 3 min time-lapse series (at 3 s acquisition frame rate) in SIM-TIRF mode was collected using 20% laser power with 100 ms exposure time to follow MreB-sw-mNeonGreen dynamics. Then, a single slice of a 3D-SIM Pal-mCherry (40% laser power, 100 ms exposure) and a brightfield reference image was acquired. Raw fluorescence images were reconstructed using Nikon Elements 5.11 acquisition software with indicated settings: MreB illumination contrast 0.8, noise suppression 0.3 and blur suppression 0.05; Pal illumination contrast 3.75, noise suppression 0.1 and blur suppression 0.5. Only reconstructed images with a quality score ≥8 and passed SIMcheck quality test 84 were used for further analysis. Subsequently, MreB time-lapse series were overlayed over the reference channels in Fiji.

Particle tracking was performed in Fiji using the TrackMate v6.0.1 plugin 85 . MreB filaments were detected using the LoG-detector with an estimated radius of 0.3 µm. Spurious spots were filtered using a quality threshold of 50. Spots were linked using a Kalman filter with an initial search radius of 0.2 µm and search radius of 0.1 µm. No frame gaps were allowed. Only tracks consisting of ≥4 continuous spots (12 s) and that travelled less than 1 µm in total distance were kept for further analysis. To analyse the nature of the displacement of each track, the mean square displacement (MSD) was calculated using the MATLAB class msdanalyzer 86 . Slopes ( α ) of the individual MSD curves were extracted using the log-log fit of the MSD and the delay time τ . As the maximum delay time of 75% of the track length was used, tracks with an \(R^2\,{{{\mathrm{for}}}}\,{{{\mathrm{log}}}}\left[ {\mathrm{MSD}} \right]\) versus log[ t ] below 0.95 indicative of a poor fit to the MSD curve were excluded from the analysis. MreB filaments engaged in active cell wall synthesis are displaced by the enzymatic activities of RodA and PBP2b 41 , 42 , 43 , hence their MSD curves display slopes of α  ≈ 2 indicative of a transported particle motion above the rate of Brownian diffusion (Extended Data Fig. 10b). MreB filaments in constricting cells, as determined by the presence of Pal-mCherry foci at the division site, were analysed by fitting a 200 nm wide region of interest to the cell division site. Directional MreB tracks were deemed to contribute to the elongation of the division site. Early and late division stages were distinguished by the presence of two separated Pal foci or a continuous fluorescent signal across the cell, respectively.

Statistical analysis

All data measurements were plotted and analysed using GraphPad Prism 9 (Version 9.3.1). In general, (log-) normal distribution was tested using Shapiro-Wilk test. For comparisons of two groups, significance was determined by two-tailed, unpaired Student’s t -test with Welch correction and F -test for variance analysis. One-way analysis of variance (ANOVA) was used for comparison of more than two groups using the recommended post-test for selected pairwise comparisons. All experiments were carried out with at least 3 independent biological replicates. P values less than 0.05 were considered statistically significant.

Reporting summary

Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Data availability

The data, plasmids and strains that support the findings of this study are available from the corresponding authors on reasonable request. Representative tomograms are deposited in EMDB: EMD-27479 (wild-type), EMD-27484 ( ftsN-∆SPOR ), EMD-27485 (∆envC and ∆envC ∆nlpD ) and EMD-27486 ( ftsL* ). Corresponding raw movie frames and stacks of tilt-series are deposited as EMPIAR-11090 (wild-type), EMPIAR-11087 ( ftsN-∆SPOR ), EMPIAR-11089 ( ∆envC and ∆envC ∆nlpD ) and EMPIAR-11088 ( ftsL* ). Source data are provided with this paper.

Code availability

Scripts used in this study are deposited on GitHub at https://github.com/NavarroVettiger/Navarro-et-al_2022 .

Silhavy, T. J., Kahne, D. & Walker, S. The bacterial cell envelope. Cold Spring Harb. Perspect. Biol. 2 , a000414 (2010).

Article   PubMed   PubMed Central   Google Scholar  

Konovalova, A., Kahne, D. E. & Silhavy, T. J. Outer membrane biogenesis. Annu. Rev. Microbiol. 71 , 539–556 (2017).

Article   CAS   PubMed   PubMed Central   Google Scholar  

Rohs, P. D. A. & Bernhardt, T. G. Growth and division of the peptidoglycan matrix. Annu. Rev. Microbiol . https://doi.org/10.1146/annurev-micro-020518-120056 (2021).

Adams, D. W. & Errington, J. Bacterial cell division: assembly, maintenance and disassembly of the Z ring. Nat. Rev. Microbiol. 7 , 642–653 (2009).

Article   CAS   PubMed   Google Scholar  

Mahone, C. R. & Goley, E. D. Bacterial cell division at a glance. J. Cell Sci. 133 , jcs237057 (2020).

Beveridge, T. J. in International Review of Cytology Vol. 72 (eds Bourne, G. H. et al.) 229–317 (Academic Press, 1981).

Matias, V. R. F. & Beveridge, T. J. Cryo-electron microscopy of cell division in Staphylococcus aureus reveals a mid-zone between nascent cross walls. Mol. Microbiol. 64 , 195–206 (2007).

Matias, V. R. F., Al-Amoudi, A., Dubochet, J. & Beveridge, T. J. Cryo-transmission electron microscopy of frozen-hydrated sections of Escherichia coli and Pseudomonas aeruginosa . J. Bacteriol. 185 , 6112–6118 (2003).

Burdett, I. D. & Murray, R. G. Electron microscope study of septum formation in Escherichia coli strains B and B-r during synchronous growth. J. Bacteriol. 119 , 1039–1056 (1974).

Burdett, I. D. & Murray, R. G. Septum formation in Escherichia coli : characterization of septal structure and the effects of antibiotics on cell division. J. Bacteriol. 119 , 303–324 (1974).

Khanna, K., López-Garrido, J., Sugie, J., Pogliano, K. & Villa, E. Asymmetric localization of the cell division machinery during Bacillus subtilis sporulation. eLife 10 , e62204 (2021).

Erickson, H. P. How bacterial cell division might cheat turgor pressure – a unified mechanism of septal division in Gram-positive and Gram-negative bacteria. BioEssays 39 , 1700045 (2017).

Article   Google Scholar  

Hayles, M. F. et al. The making of frozen-hydrated, vitreous lamellas from cells for cryo-electron microscopy. J. Struct. Biol. 172 , 180–190 (2010).

Article   PubMed   Google Scholar  

Marko, M., Hsieh, C., Schalek, R., Frank, J. & Mannella, C. Focused-ion-beam thinning of frozen-hydrated biological specimens for cryo-electron microscopy. Nat. Methods 4 , 215–217 (2007).

Medeiros, J. M. et al. Robust workflow and instrumentation for cryo-focused ion beam milling of samples for electron cryotomography. Ultramicroscopy 190 , 1–11 (2018).

Rigort, A. et al. Focused ion beam micromachining of eukaryotic cells for cryoelectron tomography. Proc. Natl Acad. Sci. USA 109 , 4449–4454 (2012).

Villa, E., Schaffer, M., Plitzko, J. M. & Baumeister, W. Opening windows into the cell: focused-ion-beam milling for cryo-electron tomography. Curr. Opin. Struct. Biol. 23 , 771–777 (2013).

de Winter, D. A. M. et al. In-situ integrity control of frozen-hydrated, vitreous lamellas prepared by the cryo-focused ion beam-scanning electron microscope. J. Struct. Biol. 183 , 11–18 (2013).

Zhang, J., Ji, G., Huang, X., Xu, W. & Sun, F. An improved cryo-FIB method for fabrication of frozen hydrated lamella. J. Struct. Biol. 194 , 218–223 (2016).

Yang, X. et al. A two-track model for the spatiotemporal coordination of bacterial septal cell wall synthesis revealed by single-molecule imaging of FtsW. Nat. Microbiol. 6 , 584–593 (2021).

Burman, L. G. & Park, J. T. Molecular model for elongation of the murein sacculus of Escherichia coli . Proc. Natl Acad. Sci. USA 81 , 1844–1848 (1984).

Taguchi, A. et al. FtsW is a peptidoglycan polymerase that is functional only in complex with its cognate penicillin-binding protein. Nat. Microbiol. 4 , 587–594 (2019).

Du, S. & Lutkenhaus, J. Assembly and activation of the Escherichia coli divisome. Mol. Microbiol. 105 , 177–187 (2017).

Liu, B., Persons, L., Lee, L. & de Boer, P. A. J. Roles for both FtsA and the FtsBLQ subcomplex in FtsN-stimulated cell constriction in Escherichia coli . Mol. Microbiol. 95 , 945–970 (2015).

Park, K.-T., Du, S. & Lutkenhaus, J. Essential role for FtsL in activation of septal peptidoglycan synthesis. mBio 11 , e03012–e03020 (2020).

Tsang, M.-J. & Bernhardt, T. G. A role for the FtsQLB complex in cytokinetic ring activation revealed by an ftsL allele that accelerates division. Mol. Microbiol. 95 , 925–944 (2015).

Marmont, L. S. & Bernhardt, T. G. A conserved subcomplex within the bacterial cytokinetic ring activates cell wall synthesis by the FtsW-FtsI synthase. Proc. Natl Acad. Sci. USA 117 , 23879–23885 (2020).

Gerding, M. A. et al. Self-enhanced accumulation of FtsN at division sites and roles for other proteins with a SPOR domain (DamX, DedD, and RlpA) in Escherichia coli cell constriction. J. Bacteriol. 191 , 7383–7401 (2009).

Ursinus, A. et al. Murein (peptidoglycan) binding property of the essential cell division protein FtsN from Escherichia coli . J. Bacteriol. 186 , 6728–6737 (2004).

Yahashiri, A., Jorgenson, M. A. & Weiss, D. S. Bacterial SPOR domains are recruited to septal peptidoglycan by binding to glycan strands that lack stem peptides. Proc. Natl Acad. Sci. USA 112 , 11347–11352 (2015).

Heidrich, C. et al. Involvement of N-acetylmuramyl-L-alanine amidases in cell separation and antibiotic-induced autolysis of Escherichia coli . Mol. Microbiol. 41 , 167–178 (2001).

Uehara, T., Parzych, K. R., Dinh, T. & Bernhardt, T. G. Daughter cell separation is controlled by cytokinetic ring-activated cell wall hydrolysis. EMBO J. 29 , 1412–1422 (2010).

Judd, E. M. et al. Distinct constrictive processes, separated in time and space, divide Caulobacter inner and outer membranes. J. Bacteriol. 187 , 6874–6882 (2005).

Uehara, T., Dinh, T. & Bernhardt, T. G. LytM-domain factors are required for daughter cell separation and rapid ampicillin-induced lysis in Escherichia coli . J. Bacteriol. 191 , 5094–5107 (2009).

Ishino, F. et al. New mutations fts-36, lts-33, and ftsW clustered in the mra region of the Escherichia coli chromosome induce thermosensitive cell growth and division. J. Bacteriol. 171 , 5523–5530 (1989).

Ueki, M., Wachi, M., Jung, H. K., Ishino, F. & Matsuhashi, M. Escherichia coli mraR gene involved in cell growth and division. J. Bacteriol. 174 , 7841–7843 (1992).

Kuru, E. et al. In situ probing of newly synthesized peptidoglycan in live bacteria with fluorescent D-amino acids. Angew. Chem. Int. Ed. 51 , 12519–12523 (2012).

Article   CAS   Google Scholar  

Hale, C. A. & de Boer, P. A. Direct binding of FtsZ to ZipA, an essential component of the septal ring structure that mediates cell division in E. coli . Cell 88 , 175–185 (1997).

Buss, J. et al. In vivo organization of the FtsZ-ring by ZapA and ZapB revealed by quantitative super-resolution microscopy. Mol. Microbiol. 89 , 1099–1120 (2013).

Squyres, G. R. et al. Single-molecule imaging reveals that Z-ring condensation is essential for cell division in Bacillus subtilis . Nat. Microbiol. 6 , 553–562 (2021).

Domínguez-Escobar, J. et al. Processive movement of MreB-associated cell wall biosynthetic complexes in bacteria. Science 333 , 225–228 (2011).

Garner, E. C. et al. Coupled, circumferential motions of the cell wall synthesis machinery and MreB filaments in B. subtilis . Science 333 , 222–225 (2011).

van Teeffelen, S. et al. The bacterial actin MreB rotates, and rotation depends on cell-wall assembly. Proc. Natl Acad. Sci. USA 108 , 15822–15827 (2011).

Dion, M. F. et al. Bacillus subtilis cell diameter is determined by the opposing actions of two distinct cell wall synthetic systems. Nat. Microbiol. 4 , 1294–1305 (2019).

Hussain, S. et al. MreB filaments align along greatest principal membrane curvature to orient cell wall synthesis. eLife 7 , e32471 (2018).

Ursell, T. S. et al. Rod-like bacterial shape is maintained by feedback between cell curvature and cytoskeletal localization. Proc. Natl Acad. Sci. USA 111 , E1025–E1034 (2014).

Truong, T. T., Vettiger, A. & Bernhardt, T. G. Cell division is antagonized by the activity of peptidoglycan endopeptidases that promote cell elongation. Mol. Microbiol. 114 , 966–978 (2020).

Paradis-Bleau, C. et al. Lipoprotein cofactors located in the outer membrane activate bacterial cell wall polymerases. Cell 143 , 1110–1120 (2010).

Typas, A. et al. Regulation of peptidoglycan synthesis by outer-membrane proteins. Cell 143 , 1097–1109 (2010).

Busiek, K. K. & Margolin, W. A role for FtsA in SPOR-independent localization of the essential Escherichia coli cell division protein FtsN. Mol. Microbiol. 92 , 1212–1226 (2014).

Lyu, Z. et al. FtsN activates septal cell wall synthesis by forming a processive complex with the septum-specific peptidoglycan synthase in E. coli. Preprint at bioRxiv https://doi.org/10.1101/2021.08.23.457437 (2021).

Begg, K. J. et al. The balance between different peptidoglycan precursors determines whether Escherichia coli cells will elongate or divide. J. Bacteriol. 172 , 6697–6703 (1990).

Lleo, M. M., Canepari, P. & Satta, G. Bacterial cell shape regulation: testing of additional predictions unique to the two-competing-sites model for peptidoglycan assembly and isolation of conditional rod-shaped mutants from some wild-type cocci. J. Bacteriol. 172 , 3758–3771 (1990).

Miller, J. H. Experiments in Molecular Genetics (Cold Spring Harbor Laboratory, 1972).

Datsenko, K. A. & Wanner, B. L. One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc. Natl Acad. Sci. USA 97 , 6640–6645 (2000).

Dubochet, J. et al. Cryo-electron microscopy of vitrified specimens. Q. Rev. Biophys. 21 , 129–228 (1988).

Wolff, G. et al. Mind the gap: micro-expansion joints drastically decrease the bending of FIB-milled cryo-lamellae. J. Struct. Biol. 208 , 107389 (2019).

Wagner, F. R. et al. Preparing samples from whole cells using focused-ion-beam milling for cryo-electron tomography. Nat. Protoc. 15 , 2041–2070 (2020).

Mastronarde, D. N. SerialEM: a program for automated tilt series acquisition on Tecnai microscopes using prediction of specimen position. Microsc. Microanal. 9 , 1182–1183 (2003).

Hagen, W. J. H., Wan, W. & Briggs, J. A. G. Implementation of a cryo-electron tomography tilt-scheme optimized for high resolution subtomogram averaging. J. Struct. Biol. 197 , 191–198 (2017).

Kremer, J. R., Mastronarde, D. N. & McIntosh, J. R. Computer visualization of three-dimensional image data using IMOD. J. Struct. Biol. 116 , 71–76 (1996).

Mastronarde, D. N. & Held, S. R. Automated tilt series alignment and tomographic reconstruction in IMOD. J. Struct. Biol. 197 , 102–113 (2017).

Xiong, Q., Morphew, M. K., Schwartz, C. L., Hoenger, A. H. & Mastronarde, D. N. CTF determination and correction for low dose tomographic tilt series. J. Struct. Biol. 168 , 378–387 (2009).

Castaño-Díez, D., Kudryashev, M., Arheit, M. & Stahlberg, H. Dynamo: a flexible, user-friendly development tool for subtomogram averaging of cryo-EM data in high-performance computing environments. J. Struct. Biol. 178 , 139–151 (2012).

Castaño-Díez, D. The Dynamo package for tomography and subtomogram averaging: components for MATLAB, GPU computing and EC2 Amazon Web Services. Acta Crystallogr. D 73 , 478–487 (2017).

Castaño-Díez, D., Kudryashev, M. & Stahlberg, H. Dynamo catalogue: geometrical tools and data management for particle picking in subtomogram averaging of cryo-electron tomograms. J. Struct. Biol. 197 , 135–144 (2017).

Navarro, P. P., Stahlberg, H. & Castaño-Díez, D. Protocols for subtomogram averaging of membrane proteins in the Dynamo software package. Front. Mol. Biosci. 5 , 82 (2018).

Koenderink, J. J. & van Doorn, A. J. Surface shape and curvature scales. Image Vis. Comput. 10 , 557–564 (1992).

Navarro, P. P. Quantitative cryo-electron tomography. Front. Mol. Biosci. 9 , 934465 (2022).

Schindelin, J. et al. Fiji: an open-source platform for biological-image analysis. Nat. Methods 9 , 676–682 (2012).

Allouche, J. et al. NNT mediates redox-dependent pigmentation via a UVB- and MITF-independent mechanism. Cell 184 , 4268–4283.e20 (2021).

Navarro, P., Scaramuzza, S., Stahlberg, H. & Castaño-Díez, D. The Dynamo software pPackage for cryo-electron tomography and subtomogram averaging. Microsc. Microanal. 26 , 3142–3145 (2020).

Pettersen, E. F. et al. UCSF Chimera–a visualization system for exploratory research and analysis. J. Comput. Chem. 25 , 1605–1612 (2004).

Buss, J. A., Peters, N. T., Xiao, J. & Bernhardt, T. G. ZapA and ZapB form an FtsZ-independent structure at midcell. Mol. Microbiol. 104 , 652–663 (2017).

Ringel, P. D., Hu, D. & Basler, M. The role of type VI secretion system effectors in target cell lysis and subsequent horizontal gene transfer. Cell Rep. 21 , 3927–3940 (2017).

Mangeol, P., Prevo, B. & Peterman, E. J. G. KymographClear and KymographDirect: two tools for the automated quantitative analysis of molecular and cellular dynamics using kymographs. Mol. Biol. Cell 27 , 1948–1957 (2016).

Reshes, G., Vanounou, S., Fishov, I. & Feingold, M. Cell shape dynamics in Escherichia coli . Biophys. J. 94 , 251–264 (2008).

Söderström, B., Chan, H., Shilling, P. J., Skoglund, U. & Daley, D. O. Spatial separation of FtsZ and FtsN during cell division. Mol. Microbiol. 107 , 387–401 (2018).

Bisson-Filho, A. W. et al. Treadmilling by FtsZ filaments drives peptidoglycan synthesis and bacterial cell division. Science 355 , 739–743 (2017).

Ursell, T. et al. Rapid, precise quantification of bacterial cellular dimensions across a genomic-scale knockout library. BMC Biol. 15 , 17 (2017).

Liang, H. et al. Metabolic labelling of the carbohydrate core in bacterial peptidoglycan and its applications. Nat. Commun. 8 , 15015 (2017).

Yang, X. et al. A two-track model for the spatiotemporal coordination of bacterial septal cell wall synthesis revealed by single-molecule imaging of FtsW. Nat. Microbiol . https://doi.org/10.1038/s41564-020-00853- 0 (2021).

Fumeaux, C. & Bernhardt, T. G. Identification of MupP as a new peptidoglycan recycling factor and antibiotic resistance determinant in Pseudomonas aeruginosa. mBio 8 , e00102–17 (2017).

Ball, G. et al. SIMcheck: a toolbox for successful super-resolution structured illumination microscopy. Sci. Rep. 5 , 15915 (2015).

Tinevez, J.-Y. et al. TrackMate: an open and extensible platform for single-particle tracking. Methods 115 , 80–90 (2017).

Tarantino, N. et al. TNF and IL-1 exhibit distinct ubiquitin requirements for inducing NEMO–IKK supramolecular structures. J. Cell Biol. 204 , 231–245 (2014).

Download references

Acknowledgements

We thank P. V. Dip, E. Brignole and A. Osherov at the MIT.nano cryo-EM facility, K. Song and C. Xu at the University of Massachusetts cryo-EM facility and R. Walsh, S. Sterling and Z. Li at the cryo-EM @ Harvard Medical School facility for providing access to the cryo-EM microscopes and for all their help, advice and maintenance of cryo-EM equipment. We also thank the MicRoN imaging core at Harvard Medical School for excellent advice on live cell imaging and maintenance of fluorescence microscopes; C. Saenz and the Microfabrication Core facility at the department of Systems Biology at Harvard Medical School for the micro-pillar design, fabrication and consultations; and T. Bartlett and E. Kuru for advice and helpful discussions on FDAA and MurNAc-alkyne labelling experiments. P.P.N. was supported by the Swiss National Science Foundation (SNF) with both Early Postdoc.Mobility P2BSP3_188112 and Postdoc.Mobility fellowships P400PB_199252. A.V. was supported by an EMBO long-term postdoctoral fellowship ALTF_89-2019 and an SNF Postdoc.Mobility fellowship P500PB_203143. C.A. was funded by Charles University with a PRIMUS grant (PRIMUS/20/SCI/015). This work was also supported by funding from the National Institutes of Health (R35GM142553 to L.H.C. and R01AI083365 to T.G.B.) and Investigator funds from the Howard Hughes Medical Institute (T.G.B.).

Author information

These authors contributed equally, and sequence was determined alphabetically: Paula P. Navarro, Andrea Vettiger.

Authors and Affiliations

Department of Molecular Biology, Massachusetts General Hospital, Boston, MA, USA

Paula P. Navarro, Virly Y. Ananda & Luke H. Chao

Department of Genetics, Blavatnik Institute, Harvard Medical School, Boston, MA, USA

Paula P. Navarro & Luke H. Chao

Department of Microbiology, Blavatnik Institute, Harvard Medical School, Boston, MA, USA

Andrea Vettiger & Thomas G. Bernhardt

MicRoN Core, Harvard Medical School, Boston, MA, USA

Paula Montero Llopis

Faculty of Mathematics and Physics, Mathematical Institute, Charles University, Prague, Czech Republic

Christoph Allolio

Howard Hughes Medical Institute, Harvard Medical School, Boston, MA, USA

Thomas G. Bernhardt

You can also search for this author in PubMed   Google Scholar

Contributions

P.P.N and A.V. conceived the project, performed experiments, and analysed and interpreted the data. P.P.N performed cryo-FIB/cryo-ET and image processing. A.V. generated mutants and performed fluorescence microscopy experiments. V.Y.A. performed 3D segmentations of cryo-ET data. P.M.L. established the SIM-TIRF workflow and assisted in data collection. C.A. contributed to cell morphology analyses. L.H.C and T.G.B provided infrastructure and scientific advice. P.P.N, A.V., L.H.C. and T.G.B. wrote the manuscript with input from all authors.

Corresponding authors

Correspondence to Thomas G. Bernhardt or Luke H. Chao .

Ethics declarations

Competing interests.

The authors declare no competing interests.

Peer review

Peer review information.

Nature Microbiology thanks Kanika Khanna and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.

Additional information

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Extended data

Extended data fig. 1 cryo-fib / cryo-et pipeline utilized in this study..

Schematic cartoons showing the steps in sample preparation for cryo-ET. In brief, bacteria are grown to OD 600  = 0.3 and applied onto an EM grid for vitrification in liquid ethane 56 . Cryo-EM grids are kept in liquid nitrogen until transfer into the cryo-FIB microscope for milling. An illustration shows the result of milling vitrified bacteria distributed onto the holey carbon film on the mesh EM grid. (a-d) Images taken from the Aquilos Thermo Fisher Scientific graphical user interface during cryo-FIB milling performance. ( a ) Target bacteria (red circles) are first identified by SEM, e - beam. Yellow box indicates region visualized in ( b ) and magenta box indicates the area visualized in (c). (b) Corresponding FIB, ion Ga + beam, view (52° with respect to the e - beam 58 ) of the targeted grid square in (a) (yellow box). Green box indicates region visualized in (d). ( c ) SEM view of the same region shown in (a) and (b) after platinum deposition and milling. Cyan box indicates obtained lamella shown in (e). ( d ) FIB view of region shown in (b) (green box) after platinum deposition and milling. Scale bars in (a-d) are indicated on each image. After milling, cryo-EM grids containing bacterial lamellae are transferred into a TEM microscope. ( e ) Low magnification TEM 2D image of the lamella shown in (c-d), cyan box. Dashed black box indicates target region for cryo-ET acquisition. Dashed white line indicates the tilt axis for cryo-ET data acquisition. ( f ) 3D slice of the cryo-electron tomogram obtained from 3D reconstruction of aligned cryo-ET tilt series acquired in (e) (dashed black box). Scale bars: e = 1000 nm; f = 200 nm. ( g ) A representative lamella from a wild-type E. coli cell imaged at indicated imaging conditions. White box highlights region for corresponding high-magnification acquisition. Scale bars = 1 µm (low magnification); 200 nm (high magnification and cryo-electron tomogram).

Extended Data Fig. 2 Distance measurements in cryo-ET data of dividing E. coli cells.

Three dimensional slices visualizing the division site during ( a ) constriction, ( b ) septation and ( c ) cytokinesis. Dashed white line indicates OM-OM distance and white bold line indicates IM-IM distance. (a.i-c.i) Measured distances in nm of IM-IM, OM-OM and the difference between these distances at (a.i) constriction (N = 7 (wt); 10 ( ftsN-∆SPOR ); 5 ( ∆envC ); 4 ( ∆nlpD ∆envC ); 8 ( ftsL* ) images), (b.i) septation (N = 5 (wt); 4 ( ftsN-∆SPOR ); 3 ( ∆envC ); 2 ( ∆nlpD ∆envC ); 3 ( ftsL* ) images), and (c.i) cytokinesis (N = 5 (wt); 4 ( ftsN-∆SPOR ); 11 ( ∆envC ); 3 ( ∆nlpD ∆envC ); 4 ( ftsL* ) images). (a.ii-b.ii) Measured distances in nm for each strain grouped. Scale bars = 200 nm. All data are expressed as mean + SEM. ( d - f ) Schematic representing the division stages color-coded at where periplasmic width was measured. Thirty euclidean distances were measured per region (see Methods), N values for each region per stage are: (d) at septum (N = 14 (wt); 16 ( ftsN-∆SPOR ); 7 ( ∆envC ); 6 ( ∆nlpD ∆envC ); 10 ( ftsL* )); at curve (N = 15 (wt); 27 ( ftsN-∆SPOR ); 13 ( ∆envC ); 8 ( ∆nlpD ∆envC ); 19 ( ftsL* )); (e) at septum (N = 10 (wt); 8 ( ftsN-∆SPOR ); 6 ( ∆envC ); 6 ( ∆nlpD ∆envC ); 6 ( ftsL* )); at curve (N = 19 (wt); 23 ( ftsN-∆SPOR ); 12 ( ∆envC ); 6 ( ∆nlpD ∆envC ); 12 ( ftsL* )); (f) at septum (N = 4 (wt); 3 ( ftsN-∆SPOR ); 12 ( ∆envC ); 3 ( ∆nlpD ∆envC ); 6 ( ftsL* )); at curve (N = 16 (wt); 12 ( ftsN-∆SPOR ); 39 ( ∆envC ); 6 ( ∆nlpD ∆envC ); 7 ( ftsL* )). All analyzed data points are displayed, bar represents mean + SD. Significance was tested using unpaired t-test with Welch correction for paired groups when data followed gaussian distribution and Mann-Whitney when data did not follow gaussian distribution and tested relative to wild-type. Significance difference among all groups compared was tested using Welch ANOVA and Brown-Forsythe when data followed gaussian distribution and Kruskal-Wallis test when data did not follow gaussian distribution. In (a.ii) difference plot, statistical significance is shown based on F-test statistics. Ns = non-significant, * = p < 0.05, ** = p < 0.01, *** = p < 0.001, **** = p < 0.0001.

Extended Data Fig. 3 Subtomogram averaging, NAD filtering and segmentation of the cell envelope of E. coli .

( a ) STA 3D structure of the cell envelope at the septum and side wall are displayed in Chimera using solid and surface rendering. 3D slices of averages are shown. 212 particles contributed to the septum average while 8072 particles from N = 5 tomograms contributed to the side wall average. Blue dots plotted on a tomogram slice represent particles that contributes to ‘side wall’ and yellow dots represent particles contributing to septum average. Scale bar = 40 nm. In the tomogram rendering, 100 pixels blocks in the cartesian axes correspond to 102.6 nm. ( b - f ) Gallery of corresponding zoom-in summed projected central slices of cryo-electron tomograms visualizing the indicated division mutants. First column shows original image, second column shows filtered image, and third column shows filtered image with segmentation layers indicating IM = green, PG = cyan and OM = magenta. A full cryo-ET gallery can be found in Extended Data Fig. 5 . A complete overview of the number of tomograms is reported in Supplementary Tables 2 , 3 . Scale bars = 100 nm.

Extended Data Fig. 4 Measuring cell envelope constriction from kymograph data.

( a ) Schematic representation of workflow for the generation of kymographs using Kymoclear and KymogrphaDirect software. Instantaneous constriction velocity for ( b ) IM (ZipA-sfGFP) and ( c ) OM (Pal-mCherry) are plotted against normalized cell width. Bold lines show second order polynomial fits as in Fig. 2e,f . N = 150 cells (wt); N = 48 ( ftsN-∆SPOR ); N = 81 ( ∆envC ) ; N = 68 ( ftsL* ) kymographs. ( d ) Additional examples of cell envelope constriction kymographs for the corresponding strains in panels b-c. ( e ) Duration of IM (left) and OM (right) constriction was derived from kymograph measurements. Data are represented as boxplots. The line represents median; error bars depict Min-Max range. The significance of differences were tested relative to wild-type by one-way ANOVA with Dunnett’s correction for multiple comparisons; ns = non-significant (p = 0.99), * = p < 0.05, *** = p < 0.001, **** = p < 0.0001; N = 150 cells (wt); N = 48 ( ftsN-∆SPOR ); N = 81 ( ∆envC ) ; N = 68 ( ftsL* ). Scale bar = 2 µm, in kymographs = 200 nm horizontal and 10 min vertical.

Extended Data Fig. 5 Cell division and polar morphology of E. coli viewed by cryo-ET.

Gallery of summed projected central slices of cryo-electron tomograms visualizing the indicated division mutants. Black arrowhead = division site; green arrowhead = envelope bulging. Dashed white box indicates corresponding zoom-in region show in Extended Data Fig. 3 . A complete overview of number of tomograms is reported in Supplementary Tables 2 , 3 . Scale bars = 200 nm.

Extended Data Fig. 6 Measuring bulk growth rates of E. coli cell division mutants analyzed in this study.

Growth curves were measured in biological triplicates by OD 600 readings in a 96-well plate reader at 30 °C. Data is represented as mean ± SD. ( a ) Untagged strains used for cryo-ET and FDAA labeling as well as ( b ) ∆murQ mutants expressing amgK and murU for MurNAc-alykyne labeling experiments were grown in LB. Cells harboring fluorescent fusion proteins for live-cell imaging of ( c ) cell envelope constriction or ( d ) MreB tracking were grown in M9 medium supplemented with 0.2% glucose and casamino acids.

Extended Data Fig. 7 A hyperactivated divisome leads to anisotropic cell envelope constriction.

( a ) Orthogonal views of XZ and XY slices of 3D cryo-electron tomograms of the indicated division mutants. Magenta and green arrowheads indicate OM and IM, respectively. 3D volumes are displayed in cartesian 3D grids with axes indicating the dimensions in pixels. For wild-type and ∆envC ∆nlpD 100 pixels = 102.6 nm, and for ftsN-∆SPOR and ftsL* 100 pixels = 110.3 nm. ( b ) Corresponding 3D surface segmentation renderings of OM (magenta) and IM (green) are shown on the right. ( c ) Schematic overview of a theoretical kymograph for an isotropic (left) and anisotropic (right) constriction of the cell envelope. Representative examples form 3 biological replicates for wild-type (left) and ftsL* (right) are provided. Scale bars: 200 nm (vertical); 5 min (horizontal). ( d ) An anisotropy score was calculated by taking the ratio of the constriction velocity from both sides of the cell. Red line (= 1) indicates a perfectly isotropic cell envelope constriction process. Data are represented as mean ± SD, Kruskal-Wallis with Dunn’s correction for multiple comparisons among all values was calculated, exact p values are shown. N = 65 (wt); 24 ( ftsL* ); 23 ( ftsN-∆SPOR ); 44 (∆envC ) constriction were analyzed ( e ) Cells were vertically immobilized using small micro pillars imprinted into agarose pads, allowing to image the cell division site along its long axis. Representative example of the cell envelope position in vertically imaged wt and ftsL* cells. Scale bar = 2 µm. Circularity was quantified using Morphometrics . Red line (circularity = 1) indicates a perfect circle. Black line indicates median. Two-way ANOVA with Sidak’s multiple comparison test; * = p < 0.05; *** = p < 0.001. N = 132 (wt); 172 ( ftsL* ) cells imaged in three biological replicates.

Extended Data Fig. 8 Cell wall synthesis and hydrolysis measurements.

( a ) New and old cell wall material were detected with Alexa488 labelled MurNAc-alykyne (yellow) or HADA (blue), respectively. ( b ) Representative images from labeling. ( c ) Label incorporation at the division site. Bars show the mean and dots show the average from 9 different images. ftsN-∆SPOR labeling example. ( d ) Rate of sPG synthesis (line = median). ( e ) Septal PG hydrolysis. Bar represents median + 95 % confidence interval; points indicate the average of three biological replicates. ( f ) Side wall labeling (line = median). ( g ) sPG and side wall synthesis ratio. ( h ) Representative YADA labeling pattern. Yellow arrow heads indicate polar label accumulation. ( i ) Average polar YADA fluorescence. Black line indicates mean, one-way ANOVA with Dunnett’s correction for multiple comparison. Significant differences are relative to wild-type; ns = non-significant (p = 0.76), p < 0.00. ( j ) Summed projected central 3D slices through tomograms of poles. Yellow arrowhead indicates enlarged periplasm. ( k ) Periplasm thickness in cryo-ET data at the side wall (green) and pole (red) (mean + SD). Thirty Euclidean distances were measured per region. One-way ANOVA (side wall) and Kruskal-Wallis test (pole); significance was tested among all groups within each region; ns = non-significant (p = 0.91), **** = p < 0.0001, N.A. = not applicable. ( l ) Labeling patterns observed for the pulse-chase. ( m ) Sum-projection of deconvolved images after HADA pulses as shown in Fig. 3b . ( n ) Expected labeling patterns for cells expressing sulA . New and old wall is labelled with HADA (blue) and YADA (yellow), respectively. ( o ) Representative images of indicated strains after HADA pulses. Mean side wall fluorescence intensity for ( p ) HADA and ( q ) YADA fit to a ( p ) Malthusian exponential function or ( q ) one phase exponential decay. Mean fluorescence intensity ± one SD is shown, N = 360 cells each. ( r ) Fluorescence intensity ( p , q ) was measured in original non-deconvolved (raw) SUM projections. Linear regression shows strong positive (R 2  = 0.87) correlation to deconvolved fluorescence intensity values. N = 1035 values. Scale bars = 2 µm (fluorescence) or 200 nm (cryo-ET).

Extended Data Fig. 9 Z-ring views during constriction in cryo-electron tomograms of E. coli .

Summed projections of 10 slices of XZ and XY views during constriction of indicated strains. Representative examples of all strains are shown. Green arrowheads indicate IM, magenta arrowheads indicate OM, red arrowheads indicate cytoskeletal ring and red asterisks indicate zones of diffuse signal. Note, Z-ring signal is weaker in ∆envC mutants due to issues with Z-ring condensation as shown by fluorescence microscopy (Fig. 4h-j ) Scale bar = 100 nm.

Extended Data Fig. 10 The balance between elongation and division affects cell morphology.

( a ) MreB-sw-mNeonGreen dynamics were followed by SIM-TRIF microscopy for 3 min at 3 s acquisitions per frame in indicated mutants. Time-lapse series was sum-projected and overlayed over a 3D-SIM Pal-mCherry and brightfield reference image. Larger fields of view are shown as compared to Fig. 5g and are representative from three biological replicates. Bar = 1 µm ( b ) Slopes of MSD curves ( α ) were analyzed following log-log fit to l og[ MSD ] versus l og[ t ] using the MATLAB class msdanalyzer 86 . Particles displaced by diffusive motion are characterized by a slope of their l og[ MSD ] = 1, while transported particles have slopes of 2 and constrained particles display slopes < 1. No significant difference (ns = non-significant, p = 0.0693; Kruskal Wallis test with Dunn’s correction for multiple comparisons) in the slopes of MSD curves were found indicating that MreB is displaced at a similar rate and manner in all strains. Box plot error bars displaying Min-Max range of values, blackline represents median. Tracks fit to l og[ MSD ] l og[ t ] with R 2  ≤ 0.95 ( c ) Cell length and ( d ) width was measured from three independent biological replicates for the indicated mutants using Morphometrics . Line represents median. Differences in significances were tested relative to wild-type using Kruskal-Wallis one-way ANOVA with Dunnett’s correction for multiple comparisons; * = p < 0.05, **** = p < 0.0001, ns = non-significant, p = 0.69. ( e ) MreB dynamics were imaged as in ( a ). Representative temporal SUM projections of MreB-mNeon trajectories were overlayed to brightfield images. Bar = 2 µm. ( f ) Directionally moving MreB tracks were filtered by MSD analysis (see Methods) and represented as boxplots (line indicating median; error bars depict Min-Max range) and normalized by cell area. Significance in each group was tested against non-filamented control (-Ara) by two-sided unpaired t-test; ** p < 0.01, ns = non-significant, wt p = 0.053, fstN-∆SPOR  = 0.253. ( g ) Constriction curvature values of wild-type cells are plotted against division site width in 566 cells. Linear regression (R 2  = 0.135) indicates the negative correlation between cell width at the division site and constriction angle.

Supplementary information

Supplementary information.

Supplementary Figs. 1–10, Tables 1–6 and Video legends 1–8.

Reporting Summary

Supplementary video 1.

Supplementary Video 1. In situ cell division of wild-type E. coli . Cryo-electron tomograms of wt E. coli . Time-lapse series were acquired at a rate of 7 fps in the compressed format m4v for visualization purposes. Green, cyan and magenta layers indicate segmented IM, PG and OM, respectively. Scale bars, 100 nm.

Supplementary Video 2

Supplementary Video 2. Fluorescence live-cell imaging of cell envelope constriction in E. coli division mutants. Three time-lapse series of each indicated E. coli mutants acquired at a 2:30 min:sec acquisition interval are shown. Bacteria were imaged at 30 °C on 1% agarose in M9 supplemented with 0.2% casamino acids and d -glucose. Fluorescence channels (Pal-mCherry, magenta; ZipA-sfGFP, green) were deconvolved. The video was rendered at 12 fps. Scale bar, 2 µm.

Supplementary Video 3

Supplementary Video 3. In situ cell division of ftsN-∆SPOR. Cryo-electron tomograms of ftsN-∆SPOR mutant. Time-lapse series were acquired at a rate of 7 fps in the compressed format m4v for visualization purposes. Green, cyan and magenta layers indicate segmented IM, PG and OM, respectively. Scale bars, 100 nm.

Supplementary Video 4

Supplementary Video 4. In situ cell division of ∆envC ∆nlpD . Cryo-electron tomograms of ∆envC ∆nlpD mutant. Time-lapse series were acquired at a rate of 7 fps in the compressed format m4v for visualization purposes. Green, cyan and magenta layers indicate segmented IM, PG and OM, respectively. Scale bars, 100 nm.

Supplementary Video 5

Supplementary Video 5. In situ cell division of ftsL* . Cryo-electron tomograms of ftsL* mutant. Time-lapse series were acquired at a rate of 7 fps in the compressed format m4v for visualization purposes. Green, cyan and magenta layers indicate segmented IM, PG and OM, respectively. Scale bars, 100 nm.

Supplementary Video 6

Supplementary Video 6. Cell wall hydrolysis contributes to Z -ring condensation. Three-dimensional maximum intensity projections rendered in Huygens (SVI) of indicated E. coli strain expressing Pal-mCherry (magenta) and ZipA-sfGFP (green) are shown. The video was rendered at 12 fps. Scale bar, 2 µm.

Supplementary Video 7

Supplementary Video 7. MreB filament increase in cells with blocked cell division. Cell division was inhibited by expressing SulA using 0.2% l -arabinose. A three-minute SIM-TIRF time-lapse series of MreB-sw-mNeonGreen (green) was overlayed over a bright field reference image. The video was rendered at 12 fps. Scale bar, 2 µm.

Supplementary Video 8

Supplementary Video 8. MreB filaments regularly pass either through or in direct proximity of the cell division site. A three-minute SIM-TIRF time-lapse series of MreB-sw-mNeonGreen (green) was overlayed over a 3D-SIM image of Pal-mCherry (magenta) and bright field reference image. On the right side, tracking results from TrackMate are overlayed. The video was rendered at 12 fps. Scale bar, 1 µm.

Source Data Fig. 1

Numerical data.

Source Data Fig. 2

Source data fig. 3, source data fig. 4, source data fig. 5, source data extended data fig. 2, source data extended data fig. 4, source data extended data fig. 6, source data extended data fig. 7, source data extended data fig. 8, source data extended data fig. 10, rights and permissions.

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this license, visit http://creativecommons.org/licenses/by/4.0/ .

Reprints and permissions

About this article

Cite this article.

Navarro, P.P., Vettiger, A., Ananda, V.Y. et al. Cell wall synthesis and remodelling dynamics determine division site architecture and cell shape in Escherichia coli . Nat Microbiol 7 , 1621–1634 (2022). https://doi.org/10.1038/s41564-022-01210-z

Download citation

Received : 08 October 2021

Accepted : 21 July 2022

Published : 12 September 2022

Issue Date : October 2022

DOI : https://doi.org/10.1038/s41564-022-01210-z

Share this article

Anyone you share the following link with will be able to read this content:

Sorry, a shareable link is not currently available for this article.

Provided by the Springer Nature SharedIt content-sharing initiative

This article is cited by

The reproduction process of gram-positive protocells.

• Dheeraj Kanaparthi
• Marko Lampe
• Tillmann Lueders

Scientific Reports (2024)

Insights into the assembly and regulation of the bacterial divisome

• Todd A. Cameron
• William Margolin

Nature Reviews Microbiology (2024)

Xanthomonas immunity proteins protect against the cis-toxic effects of their cognate T4SS effectors

• Gabriel U Oka
• Diorge P Souza
• Chuck S Farah

EMBO Reports (2024)

Terrabacteria: redefining bacterial envelope diversity, biogenesis and evolution

• Basile Beaud Benyahia
• Simonetta Gribaldo

In situ architecture of Opa1-dependent mitochondrial cristae remodeling

• Michelle Y Fry
• Paula P Navarro
• Luke H Chao

The EMBO Journal (2024)

Quick links

• Explore articles by subject
• Guide to authors
• Editorial policies

Sign up for the Nature Briefing: Microbiology newsletter — what matters in microbiology research, free to your inbox weekly.

EDITORIAL article

Editorial: bacterial cell wall structure and dynamics.

• 1 Department of Microbiology, Weill Institute for Cell and Molecular Biology, Ithaca, NY, United States
• 2 Cornell Institute of Host-Microbe Interactions and Disease, Cornell University, Ithaca, NY, United States
• 3 School of Biosciences, Institute of Microbiology and Infection, University of Birmingham, Birmingham, United Kingdom
• 4 Department of Biology, Interfaculty Institute of Microbiology and Infection Medicine Tübingen, University of Tübingen, Tübingen, Germany

Editorial on the Research Topic Bacterial Cell Wall Structure and Dynamics

The bacterial cell wall is a complex, mesh-like structure that in most bacteria is essential for maintenance of cell shape and structural integrity. Historically, the cell wall has been of intense research interest due to its necessity for most bacteria and absence from the eukaryotic realm, positioning it as an ideal target for some of our most powerful antibiotics ( Schneider and Sahl, 2010 ). In addition, bacterial cell wall fragments can have immunostimulatory and cytotoxic properties and thus play important roles in pathogenesis and disease ( Goldman et al., 1982 ; Fleming et al., 1986 ; Royet et al., 2011 ; Sorbara and Philpott, 2011 ; Jutras et al., 2019 ).

The cell wall consists mainly of peptidoglycan (PG), a mesh of polysaccharide strands (composed of a poly-[ N -acetylglucosamine (GlcNAc)- N -acetylmuramic acid (MurNAc)] backbone) cross-linked via short peptide bridges attached to the MurNAc residues ( Vollmer et al., 2008a ). PG is synthesized on the external face of the cytoplasm. Synthesis steps include cytoplasmic generation of the lipid-linked disaccharide-pentapeptide precursor lipid II, translocation of lipid II to the outside of the cell by flippases (MurJ and/or Amj); and finally assembly of the cell wall by penicillin-binding proteins (PBPs) and Shape, Elongation, Division, and Sporulation (SEDS) proteins ( Ruiz, 2008 ; Typas et al., 2011 ; Meeske et al., 2015 , 2016 ; Cho et al., 2016 ; Taguchi et al., 2019 ). The assembly process can be further subdivided into polymerization of the GlcNAc-MurNAc-pentapeptide via glycosyltransferase reactions catalyzed by class A PBPs and SEDS proteins, and crosslinking of the peptide sidestems into a tight meshwork by class A and B PBPs and L,D -transpeptidases in a not (yet) fully-understood manner ( Zhao et al., 2017 ). In addition, the PG mesh can be decorated with secondary cell wall polymers, such as wall teichoic acids (polyol-phosphate polymers) or capsule polysaccharides that are covalently attached to PG ( Rajagopal and Walker, 2017 ). In the case of mycobacteria, layers of polysaccharides and long-chain lipids are added to the PG layer, making the cell wall structure even more complex ( Jankute et al., 2015 ).

While the cell wall must be rigid enough to maintain high intracellular pressures and withstand environmental assaults, it also needs to be flexible enough to allow for cellular expansion. In addition to synthesis functions, the cell wall is thus also constantly broken down, turned over, and remodeled ( Park and Uehara, 2008 ; Reith and Mayer, 2011 ; Mayer et al., 2019 ). This is accomplished by a poorly-understood, remarkable group of enzymes that collectively can cleave and/or modify a variety of PG structures. So-called “autolysins,” for example, are a functionally diverse group of enzymes that cut PG crosslinks (endopeptidases), peptide sidestems (amidases, carboxypeptidases), or the sugar backbone (muramidases, lytic transglycosylases) ( Scheurwater et al., 2008 ; Vollmer et al., 2008b ). PG-acetyltransferases “decorate” MurNAc backbone structures with acetyl residues, imparting increased lysozyme resistance ( Moynihan and Clarke, 2011 ). L,D -transpeptidases ( Mainardi et al., 2008 ) orchestrate D -amino acid (DAA) exchange reactions that can replace terminal D -Ala residues with a variety of alternative DAAs ( Cava et al., 2011 ); this can be exploited to label PG with fluorescent compounds ( Kuru et al., 2012 ). Many of these systems fulfill important functions such as daughter cell separation, sacculus expansion during growth, insertion of macromolecular trans-envelope protein complexes, and PG recycling ( Scheurwater and Burrows, 2011 ; Vollmer, 2012 ; Johnson et al., 2013 ).

Due to its importance for bacterial survival and the many open questions concerning mechanistic details of synthesis and turnover, the cell wall remains at the center of a large number of active research programs. The last decade in particular has seen a resurgence in interest in the bacterial cell wall—a Pubmed search with the keywords “peptidoglycan synthesis bacteria” reveals a total of 7,762 publications, of which 3,532 (45%) were published in the last 10 years. This renewed interest has been fueled by novel imaging techniques (super-resolution imaging and the development of live cell wall stains) and by new revelations of processes that had been thought to be well-understood for decades. Some recent examples include the finding that the class A “penicillin-binding proteins” (aPBPs) require outer membrane cofactors for in vivo function ( Paradis-Bleau et al., 2010 ; Typas et al., 2010 ), and that RodA/FtsW possess glycosyltransferase activity ( Taguchi et al., 2019 ). At the same time, the cell wall remains a highly attractive target for antibiotic development, which has in the last decade become ever more important due to the rise in antibiotic resistance development.

In this special topic issue, we explore some new developments in the realm of bacterial cell wall biology. This collection of articles touches upon several cornerstones of PG research, with contributions focusing on the cell wall as a target for novel antibiotics, and aspects of its synthesis, turnover and modification.

The Cell Wall as a Target for Antibiotic Discovery

In a bioinformatics tour-de-force, Jukič et al. describe novel inhibitors of UppS, an isoprenyl transferase enzyme that catalyzes a critical step in the biosynthesis of the lipid carrier molecule undecaprenol pyrophosphate (UPP). UPP is essential for the translocation of the PG precursor lipid II and other extracellular polysaccharides and thus constitutes a promising target for a novel class of cell envelope antibiotics. These inhibitors were identified by virtual docking models that predicted molecule binding based on UppS crystal structures and their interaction with a known inhibitor, bisphosphonate BPH-629. This clever approach resulted in the identification of several inhibitors, one of them with μM range inhibitory activity.

Drug-resistant Mycobacterium tuberculosis strains are a major global threat that is not being adequately met with current drug discovery efforts. In their review article, Catalão et al. describe the history of peptidoglycan-targeting drugs and their use in mycobacteria. The authors provide a potential path forward by discussing recent advances such as therapies using β-lactam/β-lactamase inhibitor combinations and the use of phage endolysins for the treatment of mycobacterial infections.

Cell Wall Synthesis and Architecture

Using X-ray crystallography and liquid state NMR, Maya-Martinez et al. investigate the structure-function relationships of PBP4 of Staphylococcus aureus , a class C PBP that unexpectedly has no PG hydrolase ( D , D -peptidase) activity, but only transpeptidase activity. S. aureus is characterized by a very high degree of PG cross-linking and PBP4 apparently plays a major role in this hyper-crosslinking. The authors show transpeptidase activity of PBP4 with disaccharide peptides in vitro , producing dimeric, multimeric, and cyclic products. Structural studies with an active site mutant (S75C) revealed potential binding sites for the donor and acceptor stem peptides involved in the transpeptidation reaction.

Hottmann et al. report on peptidoglycan metabolism in the oral Gram-negative pathogen Tannerella forsythia (Phylum Bacteroidetes ). T. forsythia depends on an exogeneous supply of the cell wall sugar N -acetylmuramic acid (MurNAc), as it lacks genes generally essential for bacteria for de novo synthesis of the peptidoglycan precursor UDP-MurNAc. A pathway for the catabolism of MurNAc involving a MurNAc-6 kinase (MurK) and a MurNAc-6P hydrolase (MurQ etherase) was established in T. forsythia , which counteracts a proposed cell wall synthesis pathway that utilizes salvaged MurNAc from the medium. Accordingly, a mutant in murK exhibited increased tolerance to low external MurNAc concentrations, presumably since blocking MurNAc degradation enhances peptidoglycan precursor synthesis.

The exact in vivo architecture of PG is poorly understood. Li et al. used Atomic Force Microscopy (AFM) for a detailed study of PG architecture, particularly at the septum, in B. subtilis . Surprisingly, B. subtilis undergoes significant changes in thickness and overall cell wall architecture in different growth phases. Li et al. were also able to isolate and image septa at varying stages of completion, visualizing the PG dynamics of septal closure at high resolution.

In a thought-provoking perspective article, Vincent et al. present a hypothesis for the evolutionary origins of the unique mycobacterial cell wall through a series of horizontal gene transfers. They support their argument by observing the distribution of key cell-wall biosynthetic enzymes across the order, which suggests that the arabinogalactan components pre-date the outer membrane and virulence related lipids. In their article, the authors propose an experiment whereby the evolutionary origins of the leaflet could be tested by attempting to reconstruct the mycobacterial cell wall in an Actinobacterium that currently lacks this feature.

Cell Wall Turnover and Modification

Duchêne et al. describe new phenotypes for endopeptidase mutants in Lactobacillus plantarum . The mechanisms of regulation and physiological functions of cell wall lytic enzymes are still poorly understood, particularly in non-model organisms. L. plantarum is an ideal system to study PG hydrolase phenotypes due to its relatively small number of PG lytic enzymes (a “mere” twelve!). The authors carefully dissect the cell biological consequences of the loss of L. plantarum 's endopeptidases and assign new putative functions to these enzymes. This study thus lifts the curtain on endopeptidase function in a Gram-positive non-model organism, which is of particular importance given the high level of redundancy of PG lytic enzymes in many model bacteria, which ordinarily makes gene-phenotype association difficult.

During cell wall turnover in the Gram-positive pathogen S. aureus , the MurNAc-GlcNAc disaccharide is released from PG by the major autolysin Atl and its components eventually reused for PG biogenesis. Kluj et al. report on the fate of this disaccharide, which is taken up and is concomitantly phosphorylated by a phosphotransferase system (PTS) transporter. In order to facilitate PG recycling, the product MurNAc-6P-GlcNAc is split intracellularly by a novel phospho-glycosidase (MupG), constituting the first characterized representative of a novel class of phospho-muramidase enzymes distributed mainly within the Firmicutes bacteria.

Hager et al. report on an intriguing mode of attachment used by some bacteria (e.g., Bacillus anthracis and Paenibacillus alvei ) to bind cell-surface proteins to the cell envelope: pyruvylated secondary cell wall polymers act as high-affinity ligands for binding. In this study, the enzymatic pathway leading to the synthesis of pyruvylated disaccharide repeats, [-4-beta-GlcNAc-1,3-(4,6-Pyr)-beta-ManNAc-1-], of the P. alvei cell wall polymer was reconstituted. The reconstitution involved recombinant CsaB enzyme, catalyzing the attachment of a pyruvate to position 4 and 6 of ManNAc in the lipid-linked precursor molecule.

Devine provides a concise mini-review about the phosphate starvation regulation in the Gram-negative E. coli and the Gram-positive B. subtilis . In both organisms, phosphate limitation is sensed by the two-component system PhoPR. However, the mechanisms controlling the Pho response differ. In Bacillus subtilis , phosphate-limitation response is linked with wall teichoic acid metabolism. PhoR activity is controlled by biosynthetic intermediates of WTA metabolism, which either promotes or inhibits autokinase activity. In E. coli , phosphate is sensed directly through substrate-responsive conformational changes in a phosphate transporter.

Vermassen et al. give a comprehensive overview of the biochemistry and in vivo cleavage activity of PG lytic enzymes. This review highlights the “mix and match” approach that many cell wall lytic enzymes have undergone, combining different PG cleavage catalytic functions (e.g., lytic transglycosylase and peptidase activity) within the same enzyme.

The unique chemical nature of PG allows it to act as a potent signaling molecule. Irazoki et al. provide an overview of the process of PG release across a broad range of bacteria and PG sensing by a wide range of hosts. The authors highlight the multiplicity of systems to generate and sense bacterial PG and suggest that there is still a great deal to be learned about the sensing of these important molecules. They conclude that this field will be driven by the development and application of new analytical technologies to identify novel PG receptors.

Peptidoglycan recycling among many Gram-negative bacteria is achieved through a core pathway of degradation, recovery and recycling. In some pathogenic Neisseria , the recycling system is partially defective, which leads to an increase in the release of immunostimulatory PG fragments. In their review article, Schaub and Dillard discuss some of the differences between Neisserial PG turnover and other, more intensively studied bacteria such as E. coli . They conclude by proposing Neisseria sp. as an attractive model system for the study of cell wall growth and turnover due to their lower number of cell wall-active enzymes, variation in cell shape, and natural competence.

In addition to variations in glycan composition and stem-peptide composition, PG can also be O-acetylated at the C-6 of MurNAc, or, less frequently, GlcNAc. Sychanta et al. provide an overview of recent advances in understanding the biochemistry of O-acetyltransferase systems in Gram-positive and Gram-negative bacteria. They also discuss current efforts at understanding the impact of inhibiting these systems and address unanswered biological questions such as the source of acetate for wall modification.

Bacterial cell wall biology remains a major frontier, both in our quest to develop a profound understanding of fundamental microbiology and to discover novel compounds that may be used to treat infections caused by antibiotic resistant bacteria. We hope that this special issue further advances this frontier and inspires additional exploration—peptidoglycan is, in many ways, still as mysterious as it was 7,762 publications ago.

Author Contributions

All authors listed have made a substantial, direct and intellectual contribution to the work, and approved it for publication.

Conflict of Interest Statement

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Research in the TD lab was supported by National Institutes of Health (NIH) grants R01AI143704 and R01GM130971. PM was supported by a BBSRC David Phillips Fellowship (grant BB/S010122/1). CM acknowledges financial support by the German research foundation (DFG: grants MA2436/7, SFB766-A15, GRK1708-B2, and TRR261-A06) and the government of the state of Baden-Württemberg (MWK-Glycobiology/Glycobiotechnology).

Cava, F., de Pedro, M. A., Lam, H., Davis, B. M., and Waldor, M. K. (2011). Distinct pathways for modification of the bacterial cell wall by non-canonical D-amino acids. EMBO J. 30, 3442–3453. doi: 10.1038/emboj.2011.246

PubMed Abstract | CrossRef Full Text | Google Scholar

Cho, H., Wivagg, C. N., Kapoor, M., Barry, Z.P, Rohs, D. A., Suh, J. A., et al. (2016). Bacterial cell wall biogenesis is mediated by SEDS and PBP polymerase families functioning semi-autonomously. Nat. Microbiol. 1:16172. doi: 10.1038/nmicrobiol.2016.172

Fleming, T. J., Wallsmith, D. E., and Rosenthal, R. S. (1986). Arthropathic properties of gonococcal peptidoglycan fragments: implications for the pathogenesis of disseminated gonococcal disease. Infect. Immun. 52, 600–608.

PubMed Abstract | Google Scholar

Goldman, W. E., Klapper, D. G., and Baseman, J. B. (1982). Detection, isolation, and analysis of a released Bordetella pertussis product toxic to cultured tracheal cells. Infect. Immun. 36, 782–794.

Jankute, M., Cox, J. A., Harrison, J., and Besra, G. S. (2015). Assembly of the mycobacterial cell wall. Annu. Rev. Microbiol. 69, 405–423. doi: 10.1146/annurev-micro-091014-104121

Johnson, J. W., Fisher, J. F., and Mobashery, S. (2013). Bacterial cell-wall recycling. Ann. N. Y. Acad. Sci. 1277, 54–75. doi: 10.1111/j.1749-6632.2012.06813.x

Jutras, B. L., Lochhead, R. B., Kloos, Z. A., Biboy, J., Strle, K., Booth, C. J., et al. (2019). Borrelia burgdorferi peptidoglycan is a persistent antigen in patients with Lyme arthritis. Proc. Natl. Acad. Sci. U.S.A. 116, 13498–13507. doi: 10.1073/pnas.1904170116

Kuru, E., Hughes, H. V., Brown, P. J., Hall, E., Tekkam, S., Cava, F., et al. (2012). In situ probing of newly synthesized peptidoglycan in live bacteria with fluorescent D-amino acids. Angew. Chem. Int. Ed Engl. 51, 12519–12523. doi: 10.1002/anie.201206749

Mainardi, J. L., Villet, R., Bugg, T. D., Mayer, C., and Arthur, M. (2008). Evolution of peptidoglycan biosynthesis under the selective pressure of antibiotics in Gram-positive bacteria. FEMS Microbiol. Rev. 32, 386–408. doi: 10.1111/j.1574-6976.2007.00097.x

Mayer, C., Kluj, R. M., Muhleck, M., Walter, A., Unsleber, S., Hottmann, I., et al. (2019). Bacteria's different ways to recycle their own cell wall. Int. J. Med. Microbiol. doi: 10.1016/j.ijmm.2019.06.006. [Epub ahead of print].

Meeske, A. J., Riley, E. P., Robins, W. P., Uehara, T., Mekalanos, J. J., Kahne, D., et al. (2016). SEDS proteins are a widespread family of bacterial cell wall polymerases. Nature 537, 634–638. doi: 10.1038/nature19331

Meeske, A. J., Sham, L. T., Kimsey, H., Koo, B. M., Gross, C. A., Bernhardt, T. G., et al. (2015). MurJ and a novel lipid II flippase are required for cell wall biogenesis in Bacillus subtilis . Proc. Natl. Acad. Sci. U.S.A. 112, 6437–6442. doi: 10.1073/pnas.1504967112

Moynihan, P. J., and Clarke, A. J. (2011). O-Acetylated peptidoglycan: controlling the activity of bacterial autolysins and lytic enzymes of innate immune systems. Int. J. Biochem. Cell Biol. 43, 1655–1659. doi: 10.1016/j.biocel.2011.08.007

Paradis-Bleau, C., Markovski, M., Uehara, T., Lupoli, T. J., Walker, S., Kahne, D. E., et al. (2010). Lipoprotein cofactors located in the outer membrane activate bacterial cell wall polymerases. Cell 143, 1110–1120. doi: 10.1016/j.cell.2010.11.037

Park, J. T., and Uehara, T. (2008). How bacteria consume their own exoskeletons (turnover and recycling of cell wall peptidoglycan). Microbiol. Mol. Biol. Rev. 72, 211–227. doi: 10.1128/MMBR.00027-07

Rajagopal, M., and Walker, S. (2017). Envelope structures of gram-positive bacteria. Curr. Top. Microbiol. Immunol. 404, 1–44. doi: 10.1007/82_2015_5021

Reith, J., and Mayer, C. (2011). Peptidoglycan turnover and recycling in Gram-positive bacteria. Appl. Microbiol. Biotechnol. 92, 1–11. doi: 10.1007/s00253-011-3486-x

Royet, J., Gupta, D., and Dziarski, R. (2011). Peptidoglycan recognition proteins: modulators of the microbiome and inflammation. Nat. Rev. Immunol. 11, 837–851. doi: 10.1038/nri3089

Ruiz, N. (2008). Bioinformatics identification of MurJ (MviN) as the peptidoglycan lipid II flippase in Escherichia coli . Proc. Natl. Acad. Sci. U.S.A. 105, 15553–15557. doi: 10.1073/pnas.0808352105

Scheurwater, E., Reid, C. W., and Clarke, A. J. (2008). Lytic transglycosylases: bacterial space-making autolysins. Int. J. Biochem. Cell Biol. 40, 586–591. doi: 10.1016/j.biocel.2007.03.018

Scheurwater, E. M., and Burrows, L. L. (2011). Maintaining network security: how macromolecular structures cross the peptidoglycan layer. FEMS Microbiol. Lett. 318, 1–9. doi: 10.1111/j.1574-6968.2011.02228.x

Schneider, T., and Sahl, H. G. (2010). An oldie but a goodie—cell wall biosynthesis as antibiotic target pathway. Int. J. Med. Microbiol. 300, 161–169. doi: 10.1016/j.ijmm.2009.10.005

Sorbara, M. T., and Philpott, D. J. (2011). Peptidoglycan: a critical activator of the mammalian immune system during infection and homeostasis. Immunol. Rev. 243, 40–60. doi: 10.1111/j.1600-065X.2011.01047.x

Taguchi, A., Welsh, M. A., Marmont, L. S., Lee, W., Sjodt, M., Kruse, A. C., et al. (2019). FtsW is a peptidoglycan polymerase that is functional only in complex with its cognate penicillin-binding protein. Nat. Microbiol. 4, 587–594. doi: 10.1038/s41564-018-0345-x

Typas, A., Banzhaf, M., Gross, C. A., and Vollmer, W. (2011). From the regulation of peptidoglycan synthesis to bacterial growth and morphology. Nat. Rev. Microbiol. 10, 123–136. doi: 10.1038/nrmicro2677

Typas, A., Banzhaf, M. B., van den Berg van Saparoea, B, Verheul, J., Biboy, J., Nichols, R. J., et al. (2010). Regulation of peptidoglycan synthesis by outer-membrane proteins. Cell 143, 1097–1109. doi: 10.1016/j.cell.2010.11.038

Vollmer, W. (2012). Bacterial growth does require peptidoglycan hydrolases. Mol. Microbiol. 86, 1031–1035. doi: 10.1111/mmi.12059

Vollmer, W., Blanot, D., and de Pedro, M. A. (2008a). Peptidoglycan structure and architecture. FEMS Microbiol. Rev. 32, 149–167. doi: 10.1111/j.1574-6976.2007.00094.x

Vollmer, W., Joris, B., Charlier, P., and Foster, S. (2008b). Bacterial peptidoglycan (murein) hydrolases. FEMS Microbiol. Rev. 32, 259–286. doi: 10.1111/j.1574-6976.2007.00099.x

Zhao, H., Patel, V., Helmann, J. D., and Dorr, T. (2017). Don't let sleeping dogmas lie: new views of peptidoglycan synthesis and its regulation. Mol. Microbiol. 106, 847–860. doi: 10.1111/mmi.13853

Keywords: peptidoglycan, cell wall, autolysin, PG recycling, turnover

Citation: Dörr T, Moynihan PJ and Mayer C (2019) Editorial: Bacterial Cell Wall Structure and Dynamics. Front. Microbiol. 10:2051. doi: 10.3389/fmicb.2019.02051

Received: 25 July 2019; Accepted: 20 August 2019; Published: 04 September 2019.

Reviewed by:

Copyright © 2019 Dörr, Moynihan and Mayer. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY) . The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

*Correspondence: Christoph Mayer, christoph.mayer@uni-tuebingen.de

† ORCID: Tobias Dörr 0000-0003-3283-9161 Partick J. Moynihan 0000-0003-4182-6223 Christoph Mayer 0000-0003-4731-4851

Disclaimer: All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article or claim that may be made by its manufacturer is not guaranteed or endorsed by the publisher.

10.2 Mechanisms of Antibacterial Drugs

Learning objective.

• Describe the mechanisms of action associated with drugs that inhibit cell wall biosynthesis, protein synthesis, membrane function, nucleic acid synthesis, and metabolic pathways

An important quality for an antimicrobial drug is selective toxicity , meaning that it selectively kills or inhibits the growth of microbial targets while causing minimal or no harm to the host. Most antimicrobial drugs currently in clinical use are antibacterial because the prokaryotic cell provides a greater variety of unique targets for selective toxicity, in comparison to fungi, parasites, and viruses. Each class of antibacterial drugs has a unique mode of action (the way in which a drug affects microbes at the cellular level), and these are summarized in Figure 1 0 .4 and Table 1 0 .1 .

Inhibitors of Cell Wall Biosynthesis

Several different classes of antibacterials block steps in the biosynthesis of peptidoglycan, making cells more susceptible to osmotic lysis ( Table 1 0 .2 ). Therefore, antibacterials that target cell wall biosynthesis are bactericidal in their action. Because human cells do not make peptidoglycan, this mode of action is an excellent example of selective toxicity. Antibiotics that inhibit the cell wall biosynthesis of bacteria include the penicillins (including ampicillin, amoxicillin, and methicillin), cephalosporins, vancomycin, and bacitracin. Although it may be administered orally or intramuscularly in some circumstances, bacitracin has been shown to be nephrotoxic (damaging to the kidneys). Therefore, it is more commonly combined with neomycin and polymyxin in topical ointments such as Neosporin.

Some of these antibiotics are natural antibiotics produced by fungi or bacteria, while others are semi-synthetic, where a natural antibiotic has been chemically modified in the lab.

Spectrum of Activity

• Describe the mode of action of β-lactams.

Inhibitors of Protein Biosynthesis

The cytoplasmic ribosomes found in animal cells (80S) are structurally distinct from those found in bacterial cells (70S), making protein biosynthesis a good selective target for antibacterial drugs. Several types of protein biosynthesis inhibitors are discussed in this section and are summarized in Figure 1 0 . 5 and Table 10.3 .

Protein Synthesis Inhibitors That Bind the 30S Subunit

Aminoglycosides are large, highly polar antibacterial drugs that bind to the 30S subunit of bacterial ribosomes, impairing the proofreading ability of the ribosomal complex responsible for making cellular proteins. The aminoglycosides , which include drugs such as streptomycin, gentamicin, neomycin, and kanamycin, are potent broad-spectrum antibacterials. However, aminoglycosides have been shown to be nephrotoxic (damaging to kidney), neurotoxic (damaging to the nervous system), and ototoxic (damaging to the ear).

Another class of antibacterial compounds that bind to the 30S subunit is the tetracyclines . In contrast to aminoglycosides, these drugs are bacteriostatic and inhibit protein synthesis by blocking the association of tRNAs with the ribosome during translation. Although the tetracyclines are broad spectrum in their coverage of bacterial pathogens, side effects that can limit their use include phototoxicity, permanent discoloration of developing teeth, and liver toxicity with high doses or in patients with kidney impairment.

Protein Synthesis Inhibitors That Bind the 50S Subunit

There are several classes of antibacterial drugs that work through binding to the 50S subunit of bacterial ribosomes. Specific examples include erythromycin, azithromycin, and chloramphenicol. The first drug discovered in this category was erythromycin . It was isolated in 1952 from Streptomyces erythreus . Compared with erythromycin, azithromycin has a broader spectrum of activity, fewer side effects, and a significantly longer half-life (1.5 hours for erythromycin versus 68 hours for azithromycin) that allows for once-daily dosing and a short 3-day course of therapy (i.e., Zpac formulation) for most infections.

The drug chloramphenicol represents yet another structurally distinct class of antibacterials that also bind to the 50S ribosome, inhibiting peptide bond formation. Chloramphenicol, produced by Streptomyces venezuelae , was discovered in 1947; in 1949, it became the first broad-spectrum antibiotic that was approved by the FDA. Although it is a natural antibiotic, it is also easily synthesized and was the first antibacterial drug synthetically mass produced. As a result of its mass production, broad-spectrum coverage, and ability to penetrate into tissues efficiently, chloramphenicol was historically used to treat a wide range of infections, from meningitis to typhoid fever to conjunctivitis. Unfortunately, serious side effects, such as lethal gray baby syndrome, and suppression of bone marrow production, have limited its clinical role. Because of toxicity concerns, chloramphenicol usage in humans is now rare in the United States and is limited to severe infections unable to be treated by less toxic antibiotics. Because its side effects are much less severe in animals, it is used in veterinary medicine.

• Compare and contrast the different types of protein synthesis inhibitors.

Inhibitors of Membrane Function

A small group of antibacterials target the bacterial membrane as their mode of action ( Table 1 0 .4 ). The polymyxins are natural polypeptide antibiotics that were first discovered in 1947 as products of Bacillus polymyxa ; only polymyxin B and polymyxin E ( colistin ) have been used clinically. They are lipophilic with detergent-like properties and interact with the lipopolysaccharide component of the outer membrane of gram-negative bacteria, ultimately disrupting both their outer and inner membranes and killing the bacterial cells. Unfortunately, the membrane-targeting mechanism is not a selective toxicity, and these drugs also target and damage the membrane of cells in the kidney and nervous system when administered systemically. Because of these serious side effects and their poor absorption from the digestive tract, polymyxin B is used in over-the-counter topical antibiotic ointments (e.g., Neosporin), and oral colistin was historically used only for bowel decontamination to prevent infections originating from bowel microbes in immunocompromised patients or for those undergoing certain abdominal surgeries. The antibacterial daptomycin is a cyclic lipopeptide produced by Streptomyces roseosporus that seems to work like the polymyxins, inserting in the bacterial cell membrane and disrupting it. However, in contrast to polymyxin B and colistin, which target only gram-negative bacteria, daptomycin specifically targets gram-positive bacteria. It is typically administered intravenously and seems to be well tolerated, showing reversible toxicity in skeletal muscles.

• How do polymyxins inhibit membrane function?

Inhibitors of Nucleic Acid Synthesis

Some antibacterial drugs work by inhibiting nucleic acid synthesis ( Table 1 0 .5 ). The drug rifampin is a semisynthetic member of the rifamycin family and functions by blocking RNA polymerase activity in bacteria. The RNA polymerase enzymes in bacteria are structurally different from those in eukaryotes, providing for selective toxicity against bacterial cells. It is used for the treatment of a variety of infections, but its primary use, often in a cocktail with other antibacterial drugs, is against mycobacteria that cause tuberculosis. Despite the selectivity of its mechanism, rifampin can induce liver enzymes to increase metabolism of other drugs being administered (antagonism), leading to hepatotoxicity (liver toxicity) and negatively influencing the bioavailability and therapeutic effect of the companion drugs.

Fluoroquinolones, such as ciprofloxacin, kills bacterial cells by blocking DNA replication.

• Why do inhibitors of bacterial nucleic acid synthesis not target host cells?

Inhibitors of Metabolic Pathways

Some synthetic drugs control bacterial infections by functioning as antimetabolites , competitive inhibitors for bacterial metabolic enzymes ( Table 1 0 .6 ). The sulfonamides ( sulfa drugs ) are the oldest synthetic antibacterial agents and are structural analogues of para -aminobenzoic acid (PABA), an early intermediate in folic acid synthesis ( Figure 1 0.6 ). By inhibiting the enzyme involved in the production of dihydrofolic acid, sulfonamides block bacterial biosynthesis of folic acid and, subsequently, pyrimidines and purines required for nucleic acid synthesis. This mechanism of action provides bacteriostatic inhibition of growth against a wide spectrum of gram-positive and gram-negative pathogens. Because humans obtain folic acid from food instead of synthesizing it intracellularly, sulfonamides are selectively toxic for bacteria. However, allergic reactions to sulfa drugs are common. Another example of an antimetabolite that inhibits the folic acid synthesis pathway is trimethoprim , a synthetic antimicrobial compound ( Figure 1 0 .6 ). Trimethoprim is used in combination with the sulfa drug sulfamethoxazole to treat urinary tract infections, ear infections, and bronchitis. When used alone, each antimetabolite only decreases production of folic acid to a level where bacteriostatic inhibition of growth occurs. However, when used in combination, inhibition of both steps in the metabolic pathway decreases folic acid synthesis to a level that is lethal to the bacterial cell. Because of the importance of folic acid during fetal development, sulfa drugs and trimethoprim use should be carefully considered during early pregnancy.

• How do sulfonamides and trimethoprim selectively target bacteria?

Inhibitor of ATP Synthase

Bedaquiline, representing the synthetic antibacterial class of compounds called the diarylquinolines, uses a novel mode of action that specifically inhibits mycobacterial growth. Although the specific mechanism has yet to be elucidated, this compound appears to interfere with the function of ATP synthases, perhaps by interfering with the use of the hydrogen ion gradient for ATP synthesis by oxidative phosphorylation, leading to reduced ATP production. Due to its side effects, including hepatotoxicity and potentially lethal heart arrhythmia, its use is reserved for serious, otherwise untreatable cases of tuberculosis.

Link to Learning

To learn more about the general principles of antimicrobial therapy and bacterial modes of action, visit Michigan State University’s Antimicrobial Resistance Learning Site (https://openstax.org/l/22MSUantireslea), particularly pages 6 through 9.

Allied Health Microbiology Copyright © 2019 by Open Stax and Linda Bruslind is licensed under a Creative Commons Attribution-NonCommercial-ShareAlike 4.0 International License , except where otherwise noted.

Share This Book

• Primary Antibodies
• Secondary Antibodies
• Igs and Isotype Controls
• Antibody Panels
• ELISA and Immunoassays
• Biochemical Assays
• Cell-Based Assays
• Virus-based Tools
• Nucleic Acids
• Blood Products
• Biochemicals
• Reference Standards
• Dyes and Labeling Reagents
• Antibiotics
• Biological Buffers
• Screening Libraries
• cDNA Clones
• qPCR Gene Primers
• qPCR Gene Family Primers
• qPCR Control Gene Primers
• qPCR Primer Libraries
• Kit Components
• Conjugation Kits
• Sample Prep Kits
• Cloning and Expression Kits
• Mutagenesis Kits
• Sequencing Kits
• Staining Solutions
• Cell Culture Media
• Stabilizers
• Protein Markers
• DNA Ladders
• Transfection Reagents
• AAT Bioquest
• AdipoGen Life Sciences
• Akadeum Life Sciences
• Alithea Genomics
• Arigo Biolaboratories
• Assay Genie
• Atlas Antibodies
• Bertin Bioreagent
• Bethyl Laboratories
• Biomol GmbH
• BPS Bioscience
• Cayman Chemical
• Chimerigen Laboratories
• DIMA Biotech
• Elabscience
• ELK Biotechnology
• Expert Synthesis Solutions
• IBA Lifesciences
• ImmunoChemistry
• Indigo Biosciences
• IsoSciences
• Kingfisher Biotech
• LKT Laboratories
• LubioScience
• Navinci Diagnostics
• NewEast Biosciences
• NSJ Bioreagents
• ProNet Biotech
• RealTimePrimers
• Research Donors
• RevMAb Biosciences
• Rockland Immunochemicals
• South Bay Bio
• United States Biological
• Biomol Blog
• Applications
• Buffer Recipes
• Wall Posters
• Conference Calendar
• Molarity Calculator
• Cytokine Activity Conversion
• ELISA Analysis
• Fluorescence Spectrum Viewer

How Do Antibiotics Affect Cell Wall Synthesis?

β-Lactam antibiotics, including penicillins, cephalosporins , monobactams , and carbapenems , are distinguished by a lactam ring in their molecular structure and act by inhibiting the synthesis of the peptidoglycan layer of bacterial cell walls. They interfere with the transpeptidase, transglycosylase, and/or carboxypeptidase activities of species-specific membrane-bound penicillin-binding proteins (PBPs) that facilitate cross-linking of the cell wall components during the final stage of bacterial cell wall synthesis ( Figure 1 ). This occurs by binding to the terminal D-Ala-D-Ala in the lengthening peptidoglycan structure and rapidly acylating the active site serine of the PBP, followed by very slow deacylation, which inactivates the PBP. As the cell grows, it is not able to synthesize more cell wall to accommodate the expansion. As a result, the pressure inside the cell will push the plasma membrane out of a weak spot in the cell wall like a balloon, which will eventually rupture. Because the formation of a division furrow to create new daughter cells depends on the ability to synthesize a new cell wall, the cell is unable to pinch off the extra cytoplasmic material. An extremely vulnerable spheroplast is formed when the cell wall sheds entirely. In this form, bacteria lose control over their shape, and the duplication of much of their genetic and metabolic material further disrupts homeostasis, which leads to cell death.

Bacteria develop resistance to β-lactam antibiotics by synthesizing β-lactamase, an enzyme that attacks the β-lactam ring to inactivate the antibiotic. More than 1,000 unique β-lactamases have been described. To overcome this resistance, β-lactam antibiotics are usually given with β-lactamase inhibitors. Efflux and modification or deletion of porin function can also play a role in resistance to some β-lactams in conjunction with β-lactamase production, which results in resistance to virtually all β-lactams. Some bacteria area also able to acquire new PBPs that have low affinity for β-lactams. The best-known example of this strategy for resistance occurs with methicillin-resistant S. aureus (MRSA) that evolved a PBP with low affinity for all but a few recently developed β-lactams.

Figure 1 : Antibiotics interfere with various aspects of the synthesis of the peptidoglycan cell wall. Gram-negative cell walls consist of two layers external to the cell membrane—a thin layer of peptidoglycan and an outer membrane with porins that allow certain antibiotics but not glycopeptides to diffuse through to their site of action. Gram-positive bacteria have a thicker peptidoglycan layer but lack an extra outer membrane enabling glycopeptide access.

Glycopeptides and lipoglycopeptides are composed of glycosylated cyclic or polycyclic nonribosomal peptides that interfere with cell wall formation by forming a complex between the antibiotic and the C-terminal D-Ala-D-Ala dipeptide of the nascent peptidoglycan on the outer surface of the cell membrane of Gram-positive bacteria ( Figure 1 ). The formation of this complex prevents the transglycosylation and transpeptidation reactions that are necessary for completion of the peptidoglycan chain, resulting in an incomplete cell wall and subsequent cell death. They are too bulky to pass through the porin channels found in the outer membrane of Gram-negative bacteria. In addition to its interaction with D-Ala-D-Ala, some antibiotics in this group bind to lipid II, a cell wall precursor on the cytoplasmic side of the cell membrane that must translocate across the cell membrane to deliver and incorporate its disaccharide-pentapeptide monomer for cross-linking into peptidoglycan. This interaction results in membrane depolarization and eventual membrane disruption that leads to cell death. Since glycopeptides target the outer surface of the cell wall, they do not have to overcome a membrane barrier and do not interact with enzymes. Known glycopeptide resistance mechanisms instead are associated with structural modifications in the substrates for the enzymes that incorporate the final amino acids in the pentapeptide precursors. The most relevant modifications involve replacement of the C-terminal D-Ala with D-lactate or D-serine, resulting in D-Ala-D-Lac or D-Ala-D-Ser peptide sequences with reduced binding affinities for the antibiotic.

The phosphonic acid fosfomycin inhibits the growth of a wide variety of bacteria by targeting the enolpyruvyl transferase MurA, which catalyzes the first committed step of peptidoglycan synthesis that is essential for cell wall formation ( Figure 1 ). It enters cells through glycerol-3-phosphate and hexose-6-phosphate transporters. Deficiency in these transporters that cause increased efflux or decreased cellular uptake can lead to resistance. Specific murA gene mutations that produce an enzyme with reduced affinity for fosfomycin or increased expression of MurA that overwhelms the capacity of fosfomycin for growth inhibition are also known causes of resistance. Fosfomycin-inactivating enzymes have also been documented.

List Antibiotics

• Search Menu
• Sign in through your institution
• Advance articles
• Editor's Choice
• Awards & Prizes
• Thematic Issues
• Virtual Special Issues
• FEMS Journals
• FEMS Microbiology Ecology
• FEMS Microbiology Letters
• FEMS Yeast Research
• Pathogens and Disease
• FEMS Microbes
• Author Guidelines
• Submission Site
• Open Access
• Calls for Papers
• About FEMS Microbiology Reviews
• About the Federation of European Microbiological Societies
• Editorial Board
• Advertising and Corporate Services
• Self-Archiving Policy
• Journals on Oxford Academic
• Books on Oxford Academic

Article Contents

Introduction.

• Biosynthesis of UDP-N-acetylglucosamine
• Biosynthesis of UDP-N-acetylmuramic acid

Biosynthesis of the UDP-MurNAc-peptides

Side pathways, genetic organization, concluding remarks, acknowledgements.

• < Previous

Cytoplasmic steps of peptidoglycan biosynthesis

• Article contents
• Figures & tables
• Supplementary Data

Hélène Barreteau, Andreja Kovač, Audrey Boniface, Matej Sova, Stanislav Gobec, Didier Blanot, Cytoplasmic steps of peptidoglycan biosynthesis, FEMS Microbiology Reviews , Volume 32, Issue 2, March 2008, Pages 168–207, https://doi.org/10.1111/j.1574-6976.2008.00104.x

• Permissions Icon Permissions

The biosynthesis of bacterial cell wall peptidoglycan is a complex process that involves enzyme reactions that take place in the cytoplasm (synthesis of the nucleotide precursors) and on the inner side (synthesis of lipid-linked intermediates) and outer side (polymerization reactions) of the cytoplasmic membrane. This review deals with the cytoplasmic steps of peptidoglycan biosynthesis, which can be divided into four sets of reactions that lead to the syntheses of (1) UDP- N -acetylglucosamine from fructose 6-phosphate, (2) UDP- N -acetylmuramic acid from UDP- N -acetylglucosamine, (3) UDP- N -acetylmuramyl-pentapeptide from UDP- N -acetylmuramic acid and (4) d -glutamic acid and dipeptide d -alanyl- d -alanine. Recent data concerning the different enzymes involved are presented. Moreover, special attention is given to (1) the chemical and enzymatic synthesis of the nucleotide precursor substrates that are not commercially available and (2) the search for specific inhibitors that could act as antibacterial compounds.

Peptidoglycan (or murein) is a major component of the cell wall of almost all eubacteria. It is a complex heteropolymer that is composed of long glycan chains that are cross-linked by short peptides ( Rogers et al. , 1980 ). The glycan chains are made up of alternating N -acetylglucosamine (GlcNAc) and N -acetylmuramic acid (MurNAc) residues linked by β1→4 bonds. The d -lactoyl group of each MurNAc residue is substituted by a peptide stem with a composition most often seen as l -Ala-γ- d -Glu- meso -A 2 pm (or l -Lys)- d -Ala- d -Ala (A 2 pm, 2,6-diaminopimelic acid) in nascent peptidoglycan, the last d -Ala residue being removed in the mature macromolecule. Cross-linking of the glycan chains generally occurs between the carboxyl group of d -Ala at position 4 and the amino group of the diaminoacid at position 3, either directly or through a short peptide bridge. Minor variations in the glycan chain, the peptide stem or the peptide bridge are present in the bacterial world and are detailed in the accompanying review by Vollmer (2008) .

The biosynthesis of peptidoglycan is a complex process that involves c . 20 reactions that take place in the cytoplasm (synthesis of the nucleotide precursors) and on the inner side (synthesis of lipid-linked intermediates) and outer side (polymerization reactions) of the cytoplasmic membrane. The present review deals with the cytoplasmic steps of peptidoglycan synthesis; the subsequent steps are reviewed by Bouhss (2008) and Sauvage (2008) in this issue.

The cytoplasmic steps ( Fig. 1 ) can be divided into four sets of reactions: (1) formation of UDP-GlcNAc from fructose-6-phosphate, (2) formation of UDP-MurNAc from UDP-GlcNAc, (3) assembly of the peptide stem leading to UDP-MurNAc-pentapeptide and (4) ‘side’ or ‘annex’ pathways of synthesis of d -glutamic acid and the dipeptide d -alanyl- d -alanine. The discovery of the enzyme activities involved in these processes was the subject of the pioneering works by J. T. Park, J. L. Strominger, E. Ito, E. J. J. Lugtenberg, F. C. Neuhaus and others during the 1960s and 1970s. Their results have been summarized in previous reviews to which we refer the reader ( Rogers et al. , 1980 ; Neuhaus & Hammes, 1981 ; Ward, 1984 ; Walsh et al. , 1989 ; Bugg & Walsh, 1992 ; van Heijenoort, 2001 ; El Zoeiby, 2003a; , Katz & Caufield, 2003 ; Kotnik et al. , 2007a ). In the present review, we will focus on recent data concerning the specificities, the kinetic and catalytic mechanisms and the three-dimensional structures of these enzymes. Moreover, because the nucleotide precursors that are substrates (or products) of several of these activities are not commercially available, we will give special attention to their chemical or enzymatic synthesis. Finally, as these enzymes are targets for antibacterial compounds, we present an overview of the existing inhibitors and of the current search for new specific inhibitors.

Cytoplasmic steps of peptidoglycan biosynthesis. DA, diamino acid (generally meso -A 2 pm or l -Lys).

Biosynthesis of UDP- N -acetylglucosamine

In bacteria, UDP-GlcNAc biosynthesis from fructose-6-phosphate requires four successive enzyme activities: glucosamine-6-phosphate synthase (GlmS), phosphoglucosamine mutase (GlmM), glucosamine-1-phosphate acetyltransferase and N -acetylglucosamine-1-phosphate uridyltransferase (the latter two activities are carried by the GlmU bifunctional enzyme) ( Fig. 1 ). UDP-GlcNAc is also present in eukaryotes because GlcNAc is an important building block for major biomolecules such as chitin and glycoproteins. However, as the eukaryotic pathway of UDP-GlcNAc biosynthesis is different from the prokaryotic pathway, the latter can be considered to be a target for specific antibacterial compounds.

GlmS is an amidotransferase that takes part in the first committed step of hexosamine metabolism. It has been purified from Escherichia coli ( Badet et al. , 1987 ) and Thermus thermophilus ( Badet-Denisot et al. , 1997 ), and characterized. It is an essential and dimeric enzyme that catalyses the conversion of d -fructose-6-phosphate into d -glucosamine-6-phosphate, using l -glutamine as the nitrogen source. It follows an ordered bi–bi mechanism ( Badet et al. , 1988 ).

The GlmS monomer is composed of two structurally and functionally distinct domains ( Fig. 2 ). The N-terminal 30-kDa glutaminase domain promotes glutamine hydrolysis into glutamate and ammonia, whereas the C-terminal 40-kDa isomerase domain binds the nitrogen acceptor and uses the ammonia that is produced for the conversion of fructose-6-phosphate into glucosamine-6-phosphate. The glutamine hydrolysis reaction uses the N-terminal cysteine thiol, which forms a γ-glutamyl thioester intermediate. The ketose/aldose isomerase activity proceeds by abstraction of the C-1 pro-R hydrogen of a putative fructosimine-6-phosphate intermediate, to form a transient cis -enolamine that, upon reprotonation at the re face of the C-2 sp 2 carbon, yields glucosamine-6-phosphate ( Fig. 3 ) [see references in Badet-Denisot (1993) ].

Three-dimensional structure of the Escherichia coli GlmS monomer (PDB entry: 1jxa). The glutaminase and isomerase domains are shown in green and blue, respectively. Substrate fructose-6-phosphate is shown in magenta. The ammonia channel is indicated by red dots.

Mechanism of the ketose/aldose isomerization reaction catalysed by GlmS. From the reaction of the open form of fructose-6-phosphate and ammonia (originating from l -glutamine), a fructosimine intermediate is formed. Stereospecific abstraction of the pro-R proton of the C-1 of the intermediate gives a transient cis -enolamine that, upon reprotonation at the re face of the C-2 sp 2 carbon, yields the open form of glucosamine-6-phosphate. According to Golinelli-Pimpaneau (1989) .

The individual crystal structures of these two domains as complexes with their respective reaction products have been solved ( Obmolova et al. , 1994 ; Isupov et al. , 1996 ; Teplyakov et al. , 1998 ). As shown by its conserved N-terminal catalytic cysteine, GlmS belongs to the N-terminal nucleophile (Ntn) family and shares common structural and catalytic mechanisms with other Ntn family members. Two other invariant residues appear to have key roles in substrate recognition by anchoring the α-COO − and α-NH 3 + functions of glutamine. However, no sequence similarity has been detected between other isomerases and GlmS. Moreover, with respect to other isomerases, the catalytic residues of GlmS belong to different polypeptide chains, whereas they are usually located on only one subunit in other isomerases. The intact Escherichia coli GlmS crystal structure was solved ( Teplyakov et al. , 2001 ), thus allowing an understanding as to how the ammonia produced at the glutamine site is delivered to the sugar phosphate site: an intramolecular nitrogen channelling between the two active sites that are separated by 18 Å is involved. Both sites contribute to the formation of this channel due to several residues that are strictly conserved in the GlmS sequence ( Fig. 2 ). Moreover, the dimerization of GlmS appears to be crucial for the closure of the sugar-binding site and the formation of the channel. Recently, a study combining molecular dynamics simulations and site-directed mutagenesis allowed to define amino acid residues involved in the channelling process ( Floquet et al. , 2007a ).

The GlmS activity is inhibited by glutamine analogues that may be either naturally occurring or synthetic. Some of these possess an electrophilic functionality at the γ-position that can establish a covalent bond with the N-terminal cysteine residue located in the glutamine-binding site. Among the naturally occurring compounds, there is anticapsin, an epoxyamino acid that is liberated after hydrolysis of the dipeptide antibiotic bacilysin (tetaine) ( Kenig et al. , 1976 ) and that inactivates the enzyme; glutamine protects against inactivation ( Chmara et al. , 1984 ). Other natural inhibitors are known, like azaserine and albizziin ( Winterburn & Phelps, 1971 ). On the basis of these naturally occurring compounds, synthetic inhibitors have been studied, like 6-diazo-5-oxo- l -norleucine (DON) ( Badet et al. , 1987 ) and N 3 -fumaroyl l -2,3-diaminopropionic acid derivatives ( Chmara et al. , 1985 ; Badet et al. , 1988 ). Other glutamine analogues with low micromolar K i values have been synthesized, such as the γ-dimethylsulfonium derivative ( Walker et al. , 2000 ). Some carbohydrate compounds, like 2-amino-2-deoxy- d -glucitol-6-phosphate, have been shown to inhibit the enzyme competitively with respect to fructose-6-phosphate ( Badet-Denisot et al. , 1995 ); recently, other carbohydrate-based inhibitors have been designed as analogues of the reaction intermediates ( Bearne & Blouin, 2000 ). N -iodoacetylglucosamine-6-phosphate is an active site-directed irreversible inactivator of GlmS from Escherichia coli , and it interacts with both the sugar- and the glucosamine-binding sites ( Bearne, 1996 ). Very recently, new inhibitors of bacterial GlmS have been discovered through virtual screening; interestingly, these compounds were predicted to interact with the protein region that forms a pocket at the interface between the two enzyme monomers, which opens the way to new molecules that alter the dimerization process ( Floquet et al. , 2007b ).

GlmM is the second enzyme involved in the biosynthesis of UDP-GlcNAc, and it catalyses the interconversion of the glucosamine-6-phosphate and glucosamine-1-phosphate isomers. GlmM was first characterized and purified to near homogeneity in Escherichia coli ( Mengin-Lecreulx & van Heijenoort, 1996 ). Its amino acid sequence contains the characteristic signature of hexosephosphate mutases, including the serine residue (Ser102) for which phosphorylation is required for enzyme activity. Accordingly, GlmM is active only in its phosphorylated form, although the dephosphorylated form also exists in vivo and both forms of the enzyme can be separated by HPLC ( Jolly et al. , 1999 ). The GlmM enzymes from Staphylococcus aureus and Pseudomonas aeruginosa have also been purified ( Jolly et al. , 1997 ; Tavares et al. , 2000 ).

The reaction catalysed by GlmM follows a ping-pong bi–bi mechanism, where GlcN-1,6-diphosphate appears as an intermediate in the catalytic process, acting as both the first product and the second substrate ( Fig. 4 ). The GlmM enzyme also catalyses the interconversion of the 1-phosphate and 6-phosphate isomers of glucose, although with reduced rate constants ( Jolly et al. , 1999 ). The inactive, dephosphorylated form of GlmM undergoes an autophosphorylation reaction when it is incubated with ATP in the presence of divalent cations. The site of phosphorylation has been shown to be the aforementioned Ser102 residue ( Jolly et al. , 2000 ).

Equation of the GlmM reaction.

No crystal structure of the GlmM enzyme is available to date. However, the crystal structure of another member of the hexosephosphate mutase family, rabbit muscle phosphoglucomutase, has been reported ( Dai et al. , 1992 ). With this phosphoglucomutase, the estimated volume of the active site cleft involving the catalytic serine residue has been described as being large enough to accommodate an ATP molecule.

First identified in Bacillus subtilis , the GlmU protein was initially thought to be a GlcNAc-1-phosphate uridyltransferase that catalyses the formation of UDP-GlcNAc from GlcNAc-1-phosphate and UTP ( Hove-Jensen et al. , 1992 ; Mengin-Lecreulx & van Heijenoort, 1993 ). However, in Escherichia coli , GlmU was shown to be a bifunctional enzyme, catalysing both acetyltransfer and uridyltransfer during the transformation of GlcN-1-phosphate to UDP-GlcNAc ( Mengin-Lecreulx & van Heijenoort, 1994 ). Moreover, the order of the chemical reaction is not random but imposed by the enzyme: GlmU first catalyses acetyltransfer from AcCoA to GlcN-1-P with the release of GlcNAc-1-P, then uridyltransfer from UTP to GlcNAc-1-P in the presence of Mg 2+ , yielding inorganic pyrophosphate and UDP-GlcNAc ( Mengin-Lecreulx & van Heijenoort, 1994 ; Gehring et al. , 1996 ). However, it should be mentioned that, under certain conditions, GlmU is capable of catalysing the two reactions in the reverse order (i.e. uridyltransfer precedes acetyltransfer), but with greatly reduced kinetic parameters ( Pompeo et al. , 2001 ).

The construction of truncated forms of GlmU has shown that this bifunctional enzyme is organized in two domains that operate without substrate channelling and that are individually active ( Gehring et al. , 1996 ; Brown et al. , 1999 ; Pompeo et al. , 2001 ). However, both these activities are essential for cell viability ( Pompeo et al. , 2001 ). The sizes of the two domains appear to be roughly equivalent, each one representing about half of the 49-kDa protein ( Fig. 5 ).

Three-dimensional structure of the Escherichia coli GlmU monomer (PDB entry: 1hv9). The N- and C-terminal residues are marked with N and C, respectively. The acetyltransferase and uridyltransferase domains are shown in yellow and green, respectively. The α-helical arm connecting the two domains is shown in blue. Products CoA and UDP-GlcNAc are shown in magenta. The Co 2+ ion cocrystallized with GlmU is depicted as a black sphere.

The crystal structure of a truncated form of GlmU has shown that the two distinct domains are connected by a long α-helical arm ( Brown et al. , 1999 ). The C-terminal domain catalyses the first reaction, which is responsible for the CoA-dependent acetylation of GlcN-1-phosphate, and it shows sequence similarities with a number of acetyltransferases ( Mengin-Lecreulx & van Heijenoort, 1994 ; Raetz & Roderick, 1995 ; Gehring et al. , 1996 ). It is characterized by an imperfect, tandem hexapeptide repeat sequence motif [LIV]-[GAED]-X 2 -[STAV]-X ( Vaara, 1992 ), which folds into a left-handed β-helix (LβH) ( Raetz & Roderick, 1995 ) ( Fig. 5 ). As with most enzymes that contain an LβH structure, the GlmU acetyltransferase domain has a trimeric organization that is absolutely required for the activity. Moreover, the catalytic site is formed by complementary regions of contact between the three adjacent monomers, as confirmed by the crystal structures of the truncated and the entire GlmU ( Olsen & Roderick, 2001 ; Pompeo et al. , 2001 ). In the C-terminal domain, there are also four cysteine residues, which are apparently located near the active site. These do not appear to be directly involved in the catalytic process, but nevertheless two of them, Cys307 and Cys324, have important roles in the acetyltransferase activity ( Pompeo et al. , 1998 ). Recently, the crystal structure of the Escherichia coli GlmU acetyltransferase active site was determined in complexes with AcCoA and with CoA/GlcN-1-P ( Olsen et al. , 2007 ).

The second reaction that is catalysed by GlmU involves uridyltransfer from UDP to GlcNAc-1-phosphate and occurs at the N-terminal domain. This domain shares sequence homology with a variety of nucleotidyltransferases over residues Met1-Ala120, which are also known as nucleotide diphosphate sugar pyrophosphorylases, and has strict conservation of the L-X 2 -G-X-G-T-X-M-(X) 4 -P-K motif ( Mio et al. , 1998 ). Its crystal structure shows that the uridine-binding site is a large open pocket, bounded by two lobes. The first lobe comprises residues that interact with the nucleotide (Asn3-Val111 and His216-Asn227), and the second lobe comprises those residues that interact with the sugar moiety (Glu112-Val215) ( Brown et al. , 1999 ; Olsen & Roderick, 2001 ) ( Fig. 5 ). Contrary to the acetyltransferase domain, trimerization is not essential for expression of the uridyltransferase activity of GlmU; however, some of the interactions between the two domains appear to participate in the folding and stability of the N-terminal domain ( Pompeo et al. , 2001 ).

The crystal structure of the GlmU protein from Streptococcus pneumoniae has been solved both in its apo form and in its complex with UDP-GlcNAc and Mg 2+ ( Kostrewa et al. , 2001 ; Sulzenbacher et al. , 2001 ). Pneumococcal GlmU contains the common LβH motif with high-sequence conservation, suggesting that similar LβH motifs exist in all the GlmU structures and have important roles in the acetyltransferase activity.

GlmU acetyltransferase activity is inactivated in the presence of thiol-specific reagents, such as iodoacetamide and N-substituted maleimides ( Mengin-Lecreulx & van Heijenoort, 1994 ; Pompeo et al. , 1998 ). Recently, some thiol-specific reagents, including N -ethylmaleimide and its derivatives, have been reported to inhibit the growth of bacterial pathogens and are thereby speculated to inhibit GlmU ( Zentz et al. , 2002 ; Burton et al. , 2006 ).

Comparison of the prokaryotic and eukaryotic systems

In eukaryotes, the pathway for UDP-GlcNAc biosynthesis appears to be significantly different: acetyltransfer occurs on GlcN-6-P and not on GlcN-1-P and, most importantly, acetyltransferase and uridyltransferase activities are carried by two distinct monofunctional enzymes ( Fig. 6 ).

Formation of UDP-GlcNAc in prokaryotes (left) and eukaryotes (right).

In mammalian cells, the GlmS equivalent, known as glutamine: fructose-6-P amidotransferase (GFAT), is an insulin-regulated enzyme that controls the flux of glucose into the hexosamine pathway ( Traxinger & Marshall, 1991 ). GFAT is about 280 kDa in size, and it is composed of four subunits; it belongs to the Ntn-amidotransferase family and possesses a Cys1 residue, as does the Escherichia coli GlmS ( Huynh et al. , 2000 ). However, unlike its bacterial equivalent, GFAT is subject to allosteric regulation by UDP-GlcNAc ( Traxinger & Marshall, 1991 ) and can also be regulated by its product GlcN-6-P ( Broschat et al. , 2002 ). In mammals, the pyrophosphorylase that condenses UTP and GlcNAc-1-P has also been identified; purified to near homogeneity from pig liver extracts, it appears to be a homodimer that is composed of two 64-kDa subunits. It requires a divalent cation (Mn 2+ ) for activity and has an unusual specifity: at high concentrations, it uses UDP-GalNAc as a substrate as well as UDP-GlcNAc in the reverse direction and GalNAc-1-P as well as GlcNAc-1-P in the forward direction ( Szumilo et al. , 1996 ). Two isoforms of this enzyme, AGX1 and AGX2, have been identified in humans ( Peneff et al. , 2001b ).

In yeast such as Saccharomyces cerevisiae and Candida albicans , the hexosamine metabolism has also been well studied ( Milewski et al. , 2006 ) and the four different enzymes involved in this pathway have been shown to be essential for cell viability. The first reaction, which sees the formation of GlcN-6-P from fructose-6-P, is catalysed by GFA1 ( Watzele & Tanner, 1989 ). Like its mammalian homologue, this enzyme is a homotetramer of 80 kDa subunits that has been crystallized ( Raczynska et al. , 2007 ). Then, GlcN-6-P is N -acetylated by the GNA1 acetyltransferase to yield GlcNAc-6-P ( Mio et al. , 1999 ). The three-dimensional structure of GNA1 has been shown to be a dimer of two identical subunits ( Peneff et al. , 2001a ). GlcNAc-6-P is further isomerized into GlcNAc-1-P by the GlcNAc-phosphate mutase AGM1 ( Hofmann et al. , 1994 ). The crystal structures of this 60-kDa protein from Candida albicans have been reported, both for the apoform and for complexes with substrates and products ( Nishitani et al. , 2006 ). Finally, UDP-GlcNAc is produced from GlcNAc-1-P by a monofunctional UDP-GlcNAc pyrophosphorylase, known as UAP1, whose sequence is well conserved in the human homologue ( Mio et al. , 1998 ; Peneff et al. , 2001b ). The crystal structure of the Candida albicans enzyme has been solved recently. Unlike prokaryotic GlmU, which has a metal ion that acts as a cofactor, no metal ion has been seen in this candidal UAP1; instead, the terminal amino function of a conserved lysine residue occupies the virtual metal ion-binding site ( Maruyama et al. , 2007 ).

Biosynthesis of UDP- N -acetylmuramic acid

The first committed stage towards the creation of the peptidoglycan polymer involves the formation of UDP-MurNAc from UDP-GlcNAc by two enzymes: MurA and MurB ( Fig. 1 ). MurA (formerly known as MurZ) catalyses the first step of this transformation by transferring the enol pyruvate moiety of phospho enol pyruvate (PEP) to the 3′-hydroxyl of UDP-GlcNAc with the release of inorganic phosphate (P i ). The resulting product, UDP-GlcNAc -enol pyruvate, undergoes a reduction catalysed by MurB using one equivalent of NADPH and a solvent-derived proton. This two-electron reduction creates the lactyl ether of UDP-MurNAc.

The MurA reaction constitutes a rare biochemical process. The only other known enol pyruvyl transfer from PEP to an OH group with the concomitant release of P i occurs during the shikimic acid pathway, in the reaction catalysed by 5- enol pyruvylshikimate-3-phosphate synthase AroA ( Walsh et al. , 1996 ; Byczynski et al. , 2003 ). Interestingly, although MurA and AroA share only c . 25% sequence identity, they exhibit the same protein architecture. The MurA enzymes from Enterobacter cloacae and Escherichia coli have been overproduced and purified ( Marquardt et al. , 1992 ; Wanke et al. , 1992 ), thereafter serving for extensive mechanistic and structural studies. The MurA reaction pathway ( Fig. 7 ) follows an addition–elimination mechanism, with the formation of a noncovalently bound phospholactoyl-UDP-GlcNAc tetrahedral intermediate ( Marquardt et al. , 1993 ). Two other intermediates have been characterized: a phospholactoyl-enzyme adduct and an O -phosphothioketal intermediate, both of which are linked to the Cys115 residue ( Wanke & Amrhein, 1993 ; Brown et al. , 1994 ; Ramilo et al. , 1994 ). However, it was demonstrated that they are not essential for catalysis ( Kim et al. , 1996 ).

Mechanism of the addition–elimination reaction catalysed by MurA. X, Y, and Z stand for side-chains involved in the reaction. A proton is added to C-3 of PEP whereas the 3′-OH group of UDP-GlcNAc gets deprotonated. After formation of a tetrahedral intermediate (central formula), the elimination of phosphate results in the formation of UDP-GlcNAc-enolpyruvate. The initial proposal that Cys115 could function as Y and Z and Asp305 as X was recently questioned. In fact, Asp305 would be responsible for the final proton abstraction leading to the elimination of phosphate (Z). According to Marquardt (1993) and Eschenburg (2003 , 2005a) .

The X-ray structures of several forms of the protein have been solved, both unliganded and in a complex ( Schönbrunn, 1996 ; Skarzynski et al. , 1996 , 1998 ; Eschenburg & Schönbrunn, 2000 ; Schönbrunn, 2000a , b ) ( Fig. 8 ). Unliganded MurA appears as a two-domain protein with an unusual fold (inside-out α/β barrel) that is built up from the sixfold repetition of the same βαβαββ motif ( Fig. 8a ). Near the hinge region, there is an isoaspartyl residue that is the product of a posttranslational modification of the Asn67–Gly68 dipeptide moiety. Upon UDP-GlcNAc binding, the reaction follows an induced-fit mechanism in which the two-domain structure undergoes large conformational changes that lead to a closed form. This has been confirmed by fluorescence spectroscopy and small-angle X-ray scattering ( Schönbrunn, 1998 ). The Pro112–Pro121 loop containing Cys115 is flexible; it is solvent-exposed in the open conformation, but forms a lid around the interdomain section in the closed conformation ( Fig. 8b ).

Three-dimensional structures of MurA. (a) Open, ligand-free form of Enterobacter cloacae MurA (PDB entry: 1naw). (b) Closed form of Escherichia coli MurA complexed with UDP-GlcNAc and fosfomycin (PDB entry: 1uae). The two domains are shown in blue and green. The Pro112–Pro121 loop is shown in red. UDP-GlcNAc is shown as sticks in magenta. Cys115 (a) and covalently linked Cys115-fosfomycin (b) are depicted as ball-and-stick models in red.

The role of the amino acid residues involved in the reaction pathway ( Fig. 7 ) has been recently questioned. Cys115 was initially suggested to act as an acid–base catalyst in the addition–elimination reaction ( Skarzynski et al. , 1998 ; Krekel et al. , 2000 ). However, it seems that it is essential for product release only ( Eschenburg et al. , 2005a ). Similarly, Asp305, which was considered to be the base abstracting a proton from the 3′-hydroxyl group of UDP-GlcNAc ( Skarzynski et al. , 1996 ; Samland et al. , 2001 ), would be responsible for the final proton abstraction from the C-3 atom of the tetrahedral intermediate ( Eschenburg et al. , 2003 ). The Lys22 residue, which is strictly conserved in the MurA enzymes, was shown to be involved in the binding of PEP and to participate in the conformational change that leads to the formation of the catalytically competent enzyme complex ( Samland et al. , 1999 , 2001 ); it has been presumed recently to also accomplish the two-proton transfer required for the addition of the substrates ( Eschenburg et al. , 2003 ).

The stereochemical course of the reaction has been revised as well. Initial studies using PEP analogues had led to the conclusion that the proton addition step at C-3 of PEP proceeded at the 2- si face and that the stereochemistry of the pair of addition and elimination steps was anti/syn ( Kim et al. , 1995 ; Lees & Walsh, 1995 ; Skarzynski et al. , 1998 ). However, recent examination of the crystal structure of the D305A mutant complexed with the tetrahedral intermediate favours an addition at the re face of PEP and an anti / syn stereochemistry ( Eschenburg et al. , 2003 ).

MurA is the target of the naturally occurring broad-spectrum antibiotic fosfomycin, which forms a covalent adduct with the reactive Cys115 residue ( Marquardt et al. , 1994 ). In the MurA enzymes of species that are naturally resistant to fosfomycin, such as Mycobacterium tuberculosis ( De Smet, 1999 ) and Chlamydia trachomatis ( McCoy et al. , 2003 ), the corresponding Cys residue is changed into Asp. Escherichia coli MurA that contains the C115D mutation is enzymatically active and resistant to inactivation by fosfomycin ( Kim et al. , 1996 ).

Gram-negative bacteria have one copy of the murA gene ( Brown et al. , 1995 ), while Gram-positive bacteria have two ( murA1 and murA2 ), which have probably arisen from gene duplication ( Du et al. , 2000 ). The MurA1 and MurA2 enzymes from Streptococcus pneumoniae have been purified: their catalytic parameters are similar, and they are both inhibited by fosfomycin ( Du et al. , 2000 ).

It has been reported recently that UDP-MurNAc tightly binds to and inhibits Escherichia coli MurA; a possible role of the nucleotide in the regulation of peptidoglycan biosynthesis has been inferred ( Mizyed et al. , 2005 ).

MurB from Escherichia coli has been overproduced and purified. It is a 38-kDa protein that contains a stoichiometric amount of bound FAD ( Benson et al. , 1993 ; Tayeh et al. , 1995 ). It follows a ping-pong bi–bi mechanism, with weak and strong substrate inhibition by NADPH and UDP-GlcNAc- enol pyruvate, respectively. It is activated by cations, such as K + , NH 4 + and Rb + ( Dhalla et al. , 1995 ).

The reaction catalysed by MurB involves two half-reactions in which FAD serves as the redox intermediate ( Fig. 9 ). The first half-reaction is the reduction of FAD to FADH 2 by NADPH. This starts with the binding of NADPH to MurB and the transfer of the 4- pro - S hydrogen of NADPH to N-5 of the enzyme-bound flavin. The release of NADP + is followed by the binding of UDP-GlcNAc- enol pyruvate. The second half-reaction is the reduction of the vinylic enol ether by FADH 2 . Hydride transfer from the reduced flavin to C-3 of the enol pyruvyl moiety of the nucleotide substrate generates a carbanionic intermediate that is then protonated at C-2 by a solvent-equilibrated proton ( Benson et al. , 1993 , 1997b ). The stereochemistry of the reduction has been studied through the use of UDP-GlcNAc-( E )- enol butyrate as a mechanistic probe ( Lees et al. , 1996 ).

Reaction mechanism of the MurB enzyme from Escherichia coli . (a) The first half-reaction consists in the transfer of the 4- pro-S hydride from NADPH to N-5 of Enz-FAD. (b) The second half-reaction consists in the transfer of this hydride from Enz-FADH 2 to C-3 of the enol pyruvyl moiety of UDP-GlcNAc- enol pyruvate, followed by quenching at C-2 by a solvent-exchangeable proton, to yield the d -configuration of the lactyl ether product. B 1 H and B 2 H stand for side-chains (Glu325 and/or Arg 159) stabilizing by hydrogen bonding the carbanion intermediate as an enol. B 3 H stands for is a general acid catalyst (Ser229) serving as the proton donor to quench the carbanion/enol intermediate and deliver the proton at C-2. From Benson (1993 , 1995 , 1997a , b) .

The crystal structures of unliganded MurB and of its dead-end complex with UDP-GlcNAc- enol pyruvate have been solved ( Benson et al. , 1995 , 1996 , 1997a ) ( Fig. 10 ). The protein is composed of three domains: domains 1 and 2 mediate FAD binding, and domain 3 mediates the binding of its substrates. The binding of UDP-GlcNAc- enol pyruvate induces a substantial movement of domain 3. With the perdeuterated, 13 C/ 15 N-labelled MurB studied by NMR spectroscopy ( Farmer et al. , 1996 ; Constantine et al. , 1997 ), it was deduced that NADP + binds in the same pocket as UDP-GlcNAc- enol pyruvate, inducing structural changes, and that NADPH transfers a hydride to the si face of the FAD isoalloxazine ring.

Three-dimensional structures of MurB. (a) Structure of Escherichia coli MurB (type I) (PDB entry: 1mbt). (b) Structure of Staphylococcus aureus MurB (type II) (PDB entry: 1hsk). The three domains are shown in yellow, blue, and green. FAD is shown in magenta. Structural elements that are absent in type II MurB are shown in red.

Comparison of the X-ray structures of Escherichia coli and Staphylococcus aureus MurB enzymes has revealed notable distinctions with respect to their structural elements. Escherichia coli MurB, classified as type I, contains a Tyr loop and a split βαββ fold ( Fig. 10a ), whereas Staphylococcus aureus MurB, classified as type II, lacks these secondary elements ( Fig. 10b ). This has consequences on the mode of substrate binding ( Benson et al. , 2001 ). Recently, a type II MurB enzyme ( Thermus caldophilus ) was crystallized in the presence of UDP-GlcNAc- enol pyruvate. X-ray data and sequence alignments allowed the definition of two subtypes, type II-a (e.g. Staphylococcus aureus and Bacillus subtilis ) and type II-b (e.g. Thermus caldophilus and Chlamydia pneumoniae ), that contain a serine or a cysteine residue, respectively, as a proton donor to quench the carbanionic intermediate ( Kim MK, 2007 ). Six other conserved amino acids were also shown by site-directed mutagenesis to be essential for MurB activity ( Nishida et al. , 2006 ).

Inhibitors of MurA and MurB

As already mentioned, the most well-known inhibitor of MurA is fosfomycin 1 ( Fig. 11 ), an epoxide compound that reacts with the Cys115 residue. A number of novel inhibitors of MurA have been discovered recently through various high-throughput screening efforts in the pharmaceutical industry ( Dai et al. , 2002 ; DeVito et al. , 2002 ; Labaudiniere et al. , 2005 ). Three noncovalent inhibitors show submicromolar IC 50 values (IC 50 =0.2–0.9 μM); however, they also show nonspecific inhibition of DNA, RNA and protein biosynthesis ( Baum et al. , 2001 ). In a whole-cell peptidoglycan synthesis assay, two new inhibitors were identified: a derivative of diarylmethane and a substituted imidazole ( Barbosa et al. , 2002 ). Sesquiterpene lactones [e.g. 2 : IC 50 =10.3 μM ( Pseudomonas aeruginosa ) and 16.7 μM ( Escherichia coli )] were shown to alkylate the thiol group of Cys115 of MurA from Pseudomonas aeruginosa and Escherichia coli and thereby to act as irreversible inhibitors ( Bachelier et al. , 2006 ). A dodecapeptide inhibitor of MurA from Pseudomonas aeruginosa (IC 50 =200 μM) was selected by phage display and appeared to be competitive with respect to UDP-GlcNAc ( Molina-López, 2006 ).

Formulae of inhibitors of MurA and MurB.

The fluorescent dye 8-anilino-1-naphthalene sulfonate 3 binds to the solvent-exposed region of MurA from Enterobacter cloacae , and its X-ray cocrystal structure provides the basis for an alternative approach in the design of new inhibitors ( Schönbrunn, 2000a ). Thus, 5-sulfonoxy-anthranilic acid derivatives that were obtained by high-throughput screening bind to the same region of MurA and obstruct its transition from the open to the closed forms ( Eschenburg et al. , 2005b ).

Tri-substituted thiazolidinones [e.g. 4 : IC 50 ( Escherichia coli )=7.7 μM] were the first small-molecule inhibitors of MurB; they were designed to mimic the diphosphate moiety of UDP-GlcNAc- enol pyruvate and prepared by a parallel synthesis approach ( Andres et al. , 2000 ). As their bioisosteric replacement, a series of imidazolinone analogues were synthesized and found to possess potent MurB inhibitory activity (the best IC 50 values for Escherichia coli ranged from 12 to 40 μM) as well as promising antibacterial activity against Staphylococcus aureus (MIC values: 2–4 μg mL −1 ) ( Bronson et al. , 2003 ). Two inhibitors of Staphylococcus aureus MurB, with K d values in the submicromolar range (0.19 and 0.14 μM), were discovered by high-throughput screening ( Sarver et al. , 2002 ). 4-Alkyl and 4,4′-dialkyl 1,2-bis(4-chlorophenyl)pyrazolidine-3,5-dione derivatives were found to inhibit MurA (the best IC 50 values for Escherichia coli ranged from 9.8 to 50 μM) and MurB (the best IC 50 values for Escherichia coli and Staphylococcus aureus ranged from 5.1 to 50 μM) ( Kutterer et al. , 2005 ). In addition, four structurally related 3,5-dioxopyrazolidines inhibited MurB with IC 50 values between 4 and 35 μM, and also MurA and MurC to weaker extents. The crystal structure of a complex with compound 5 indicated that the 3,5-dioxopyrazolidine core occupies the same region of the MurB active site as the N -acetyl group of the substrate ( Yang et al. , 2006 ). Antibacterial activity against Gram-positive bacteria was seen for many inhibitors from the pyrazolidine-3,5-dione series; however, when tested in the presence of bovine serum albumin (BSA), the activity was lost, indicating high protein-binding properties of these compounds ( Kutterer et al. , 2005 ; Yang et al. , 2006 ). A similar decline in antibacterial activity due to the presence of BSA was also seen for thiazolyl urea and carbamate derivatives [e.g. 6 : IC 50 ( Staphylococcus aureus )=19 μg mL −1 ], which were good inhibitors of MurA and MurB as well as of the growth of some Gram-positive bacteria ( Francisco et al. , 2004 ).

Generality of the Mur ligases

The stepwise assembly of the peptide stem of peptidoglycan is ensured by a series of four essential enzymes, known as the Mur ligases (MurC, D, E and F). These provide for additions of l -alanine (MurC), d -glutamic acid (MurD), a diamino acid, generally meso -diaminopimelic acid or l -lysine (MurE) and dipeptide d -Ala- d -Ala (MurF) onto the d -lactoyl group of UDP-MurNAc ( Fig. 1 ). A fifth enzyme, Mpl, adds the tripeptide l -Ala-γ- d -Glu- meso -A 2 pm directly onto UDP-MurNAc during peptidoglycan recycling. The Mur ligases catalyse the formation of an amide or a peptide bond with simultaneous formation of ADP and P i from ATP. A divalent cation, Mg 2+ or Mn 2+ , is essential for the reaction. Their mechanism of action has been studied through biochemical experiments, amino acid sequence examination, site-directed mutagenesis, assays of transition-state analogue inhibitors and X-ray structure determination. These studies have shown that the Mur ligases share three characteristics:

They have the same reaction mechanism, which consists first in the activation of the carboxyl group of the UDP-precursor by ATP, generating an acyl phosphate intermediate and ADP; the acyl phosphate then undergoes the nucleophilic attack of the amino group of the condensing amino acid (or dipeptide), leading to the formation of a high-energy tetrahedral intermediate, which eventually breaks down into amide or peptide and P i ( Fig. 12 ) [see references in Bouhss (2002) ].

They have a series of six invariant residues in addition to an ATP-binding consensus sequence. This finding led to the definition of the Mur ligases as a new family of enzymes ( Bouhss et al. , 1997 , 1999b; , Eveland et al. , 1997 ). Three other enzymes that are not related to peptidoglycan biosynthesis also belong to this family: the folylpoly-γ- l -glutamate synthetase FolC ( Sheng et al. , 2000 ), the C-terminal region of cyanophycin synthetase CphA ( Ziegler et al. , 1998 ) and the poly-γ-glutamate synthetase CapB from Bacilli ( Candela & Fouet, 2006 ).

They have the same three-dimensional structures in three domains, as seen by crystallographic studies ( Fig. 13 ). The N-terminal domain is involved in the binding of the UDP-precursor, the central domain in the binding of ATP, and the C-terminal domain in the binding of the amino acid or dipeptide. Whereas the topologies of the central and C-terminal domains are similar among the Mur ligases, that of the N-terminal domain shows differences, with MurC and MurD on the one hand, and MurE and MurF on the other hand. These differences are related to the lengths of the UDP-precursor substrates. These enzymes exist in ‘closed’ and ‘open’ conformations. The closure of the conformation is thought to be provoked by ligand binding. The description and the comparison of the three-dimensional structures of the Mur ligases has been reviewed excellently by Smith (2006) .

Reaction mechanism of the Mur ligases. First, the reaction of the carboxyl group of the UDP-precursor (R-COO − ) with ATP generates an acyl phosphate intermediate and ADP. Then, the acyl phosphate undergoes the nucleophilic attack of the amino group of the amino acid or dipeptide (R′–NH 2 ) to form a tetrahedral transition state, which eventually breaks down into amide or peptide (R–CO–NH–R′) and P i .

Three-dimensional structures of the Mur ligases. (a) Open, ligand-free form of Haemophilus influenzae MurC (PDB entry: 1gqq). (b) Closed form of Escherichia coli MurD complexed with UDP-MurNAc- l -Ala, ADP and Mg 2+ (PDB entry: 2uag). (c) Closed form of Escherichia coli MurE complexed with UDP-MurNAc- l -Ala-γ- d -Glu- meso -A 2 pm (PDB entry: 1e8c). (d) Closed form of Streptococcus pneumoniae MurF complexed with a sulfonamide inhibitor (compound 14 in Fig. 15 ) (PDB entry: 2am1). The N-terminal, central and C-terminal domains are shown in green, blue and yellow, respectively. Ligands are shown in magenta. Mg 2+ ions in (b) are depicted as black spheres. The black arrows in (a) show the direction of movement of the N- and C-terminal domains upon going from the open to the closed form.

The MurC ligase adds the first amino acid of the peptide stem. In most bacterial species, this amino acid is l -alanine; in rare cases, glycine or l -serine is added instead ( Schleifer & Kandler, 1972 ). MurC from Escherichia coli ( Liger et al. , 1995 ; Falk et al. , 1996 ; Gubler et al. , 1996 ), Mycobacterium tuberculosis and Mycobacterium leprae ( Mahapatra et al. , 2000 ), Pseudomonas aeruginosa ( El Zoeiby, 2000 ) and Chlamydia trachomatis ( Hesse et al. , 2003 ) have been purified and characterized. The enzymatic properties of MurC from Escherichia coli have been studied extensively. Its preferred substrate is l -Ala, although Gly and l -Ser, as well as several compounds that are structurally related to l -Ala, can be added with lower efficiencies ( Liger et al. , 1991 , 1995 ; Emanuele et al. , 1996 ). The stereospecificity is strict: d -Ala is not a substrate ( Liger et al. , 1995 ). A sequential, ordered kinetic mechanism has been demonstrated, with ATP binding first, UDP-MurNAc second and l -Ala third ( Emanuele et al. , 1997 ). The reaction is reversible and the exchange reaction is phosphate- and ADP-dependent ( Liger et al. , 1996 ). The existence of the acyl phosphate intermediate has been inferred from several studies, namely isotope exchange ( Falk et al. , 1996 ), rapid kinetics ( Emanuele et al. , 1997 ), radioactive labelling ( Liger et al. , 1996 ) and chemical trapping ( Bouhss et al. , 2002 ). Although MurC activity is dependent on the presence of a reducing agent, site-directed mutagenesis of the two cysteine residues present in the active site of MurC has shown that these two are not essential ( Nosal et al. , 1998 ). In solution, the Escherichia coli enzyme is present as an equilibrium between monomeric and dimeric forms; however, this has no influence on its specific activity ( Jin et al. , 1996 ).

The specificities of the MurC enzymes from species other than Escherichia coli have also been studied. Interestingly, MurC from Mycobacterium tuberculosis and Mycobacterium leprae have the same in vitro specificity patterns towards l -Ala and Gly; however, the amino acid found in the first position of the peptide stem is different ( l -Ala for the former and Gly for the latter). This appears to be due to growth conditions ( Mahapatra et al. , 2000 ). Another interesting case is that of Chlamydia trachomatis , which contains a bifunctional protein with a MurC domain and a Ddl domain ( Chopra et al. , 1998 ). The MurC domain has been purified alone ( Hesse et al. , 2003 ): it adds l -Ala, l -Ser and Gly with similar efficiencies, thereby preventing the deduction of the nature of the first amino acid of the putative chlamydial peptidoglycan ( Moulder, 1993 ). The entire MurC-Ddl fusion protein has been purified recently ( McCoy & Maurelli, 2005 ) (see ‘Formation of d -Ala- d -Ala’).

The crystal structures of the MurC enzymes from Haemophilus influenzae ( Mol et al. , 2003 ) ( Fig. 13a ), Thermotoga maritima ( Spraggon et al. , 2004 ) and Escherichia coli ( Deva et al. , 2006 ) have been solved. Unexpectedly, in the apo structures of these last two enzymes, the N-terminal and central domains were seen to be in their closed conformation, suggesting that in its unliganded form MurC may exist in different conformations in solution ( Smith, 2006 ).

The second amino-acid residue of the peptide stem is in most species d -glutamic acid ( Schleifer & Kandler, 1972 ). The few variations encountered ( d -isoglutamine, threo -3-hydroxyglutamic acid) are due to modifications at a latter stage in the biosynthesis, and thus d -Glu can be said to be the amino acid substrate of MurD in all species ( Vollmer et al. , 2008 ). The enzymes from Escherichia coli ( Pratviel-Sosa et al. , 1991 ; Auger et al. , 1998 ), Staphylococcus aureus , Haemophilus influenzae and Enterococcus faecalis ( Walsh et al. , 1999 ) have been purified and characterized. Here again, the specificity of the enzyme from Escherichia coli has been studied in detail ( Pratviel-Sosa et al. , 1994 ). An almost exclusive preference for d -Glu was seen. Only a few closely related derivatives (homocysteic acid, 3- or 4-methyl- d -Glu, cyclopentane or cyclohexane analogues of d -Glu) were fairly good substrates. l -Glu is not a substrate ( Pratviel-Sosa et al. , 1994 ); however, an N -sulfonyl derivative of l -Glu was shown to be a competitive inhibitor towards d -Glu ( Kotnik et al. , 2007b ). Among the four Mur ligases, MurD differs by its low specificity towards the UDP-precursor: whereas the UMP moiety is essential for the three other ligases, 1-phospho-MurNAc- l -Ala is a substrate for MurD ( Michaud et al. , 1987 ). The reaction is reversible; however, in contrast to MurC, the exchange reaction is not ADP-dependent ( Vaganay et al. , 1996 ). Chemical trapping experiments have established the existence of the acyl phosphate intermediate ( Bouhss et al. , 2002 ), and the tight binding of phosphinate analogues of the high-energy tetrahedral intermediate has strongly suggested its occurrence ( Tanner et al. , 1996 ; Gegnas et al. , 1998 ). MurD is capable of producing adenosine 5′-tetraphosphate, which originates from a reaction of the acyl phosphate with ATP ( Bouhss et al. , 1999a ); among the Mur ligase family, only FolC appears to share this property ( Dementin et al. , 2001 ; Sun et al. , 2001 ). MurD from other species have been characterized. Differences between Gram-negative and Gram-positive bacteria regarding substrate inhibition by UDP-MurNAc- l -Ala and effects of monovalent ions have been seen. These differences have been interpreted in terms of regulation of peptidoglycan synthesis ( Walsh et al. , 1999 ).

The first crystal structure of a Mur ligase that was solved was that of Escherichia coli MurD complexed with UDP-MurNAc- l -Ala ( Bertrand et al. , 1997 ). Other structures (complexes with products, metal ions or inhibitors, ‘open’ forms) were then reported ( Bertrand et al. , 1999 , 2001 ; Kotnik et al. , 2007b ), with strong structural similarities between MurD and FolC being seen ( Sheng et al. , 2000 ; Bertrand et al. , 2001 ). The MurD molecule contains two magnesium ions ( Fig. 13b ): the ‘classical’ one (Mg1) involved in ATP binding, and a second one (Mg2) involved in acyl phosphate formation. Mg2 is coordinated with two water molecules that are hydrogen bonded to a carbamoylated lysine residue ( Bertrand et al. , 1999 ). The importance of this carbamoyl group, which is also present in MurE and MurF, has been investigated by chemical rescue experiments ( Dementin et al. , 2001 ). Recently, a targeted molecular dynamics study has increased our understanding of the substrate binding and domain closure processes ( Perdih et al. , 2007 ).

The third amino acid of the peptide stem is generally either meso -A 2 pm (most Gram-negative bacteria and Bacilli) or l -lysine (most Gram-positive bacteria), although in certain species, other amino acids are encountered ( l -ornithine, ll -A 2 pm, meso -lanthionine, l -diaminobutyric acid, l -homoserine, for example) ( Schleifer & Kandler, 1972 ). In most cases, the MurE enzyme is highly specific for the relevant amino acid, incorporation of a ‘wrong’ amino acid (e.g. l -Lys in Escherichia coli ) leading to cell lysis ( Mengin-Lecreulx et al. , 1999 ). Bacillus sphaericus possesses two MurE enzymes: one that adds l -Lys that is active during vegetative growth, and one that adds meso -A 2 pm that is active during spore cortex formation ( Anwar & Vlaovic, 1986 ). As for the second amino acid, certain variants of the third amino acid (amidated meso -A 2 pm, acetylated diaminobutyric acid) necessitate a subsequent enzymatic activity ( Vollmer et al. , 2008 ).

The l -Lys-adding enzyme from Bacillus sphaericus and the meso -A 2 pm-adding enzyme from Escherichia coli are strongly activated by phosphate, a product of the reaction ( Anwar & Vlaovic, 1986 ; Michaud et al. , 1990 ). Previous studies on MurE from Escherichia coli have shown that a few analogues of meso -A 2 pm ( ll -A 2 pm, lanthionine and cystathionine) can be accepted as substrates either in vitro or in genetically engineered cells ( Mengin-Lecreulx et al. , 1988 , 1994 ; Michaud et al. , 1990 ; Richaud et al. , 1993 ; Auger et al. , 1996 ). The specificities of the MurE enzymes from a Gram-negative ( Escherichia coli ) and a Gram-positive ( Staphylococcus aureus ) species towards the amino acid substrate have been compared recently ( Boniface, 2007 ). While the Escherichia coli enzyme has a very weak l -Lys-adding activity, the Staphylococcus aureus MurE is totally unable to add meso -A 2 pm. Furthermore, Escherichia coli MurE does not accept l -Orn as a substrate, contrary to the staphylococcal enzyme, which has a weak l -Orn-adding activity.

In at least two species, the MurE enzyme appears to be devoid of strict specificity. MurE from Bifidobacterium globosum can incorporate two amino acids indifferently, l -Lys and l -Orn, which are both retrieved in peptidoglycan ( Hammes et al. , 1977 ). MurE from Thermotoga maritima , a Gram-negative species for which peptidoglycan contains similar proportions of both enantiomers of lysine, but no meso -A 2 pm ( Huber et al. , 1986 ), can add l -Lys, d -Lys and meso -A 2 pm in vitro with comparable efficiencies ( Boniface et al. , 2006 ). In the UDP-MurNAc-tripeptide products, the d -Glu- l -Lys bond has the conventional γ→α arrangement; however, d -Lys is acylated on its ɛ-amino group, leading to the synthesis of a new nucleotide, UDP-MurNAc- l -Ala- d -Glu(γ→ɛ) d -Lys. The absence of meso -A 2 pm in Thermotoga maritima peptidoglycan is explained by its very low intracellular pool ( Boniface et al. , 2006 ).

The X-ray structure of Escherichia coli MurE in complex with its product, UDP-MurNAc- l -Ala-γ- d -Glu- meso -A 2 pm, has been solved ( Gordon et al. , 2001 ) ( Fig. 13c ). A binding pocket for the distal (nonreacting) site of meso -A 2 pm is seen ( Fig. 14a ). Sequence alignments have revealed consensus sequences in the binding pockets of meso -A 2 pm and l -Lys: DNPR and D(D,N)P(N,A), respectively ( Dementin, 2001 ; Gordon et al. , 2001 ) ( Fig. 14b ). The main difference between these consensus sequences is the arginine residue, which is H-bonded with the carboxyl group of the distal site of meso -A 2 pm ( Gordon et al. , 2001 ) ( Fig. 14a ). In the Thermotoga maritima enzyme, the consensus sequence (DDPR) is undoubtedly a meso -A 2 pm-adding one ( Fig. 14b ), thereby explaining the ‘upside-down’ binding of d -lysine. Interestingly, MurE from Escherichia coli produces UDP-MurNAc- l -Ala- d -Glu(γ→ɛ) d -Lys in vitro , although at a low rate ( Mengin-Lecreulx et al. , 1994 ; Boniface et al. , 2006 ). The determination of the three-dimensional structure of an l -Lys-binding pocket would be of great interest; however, despite attempts with MurE from Streptococcus pneumoniae ( Blewett et al. , 2004 ) and Staphylococcus aureus ( Boniface et al. , 2007 ), no data are currently available.

Binding pocket for the distal site of meso -A 2 pm in MurE enzymes. (a) Interactions of the A 2 pm moiety of UDP-MurNAc-tripeptide with Escherichia coli MurE (PDB entry: 1e8c); Only side chains involved in hydrogen bonds (dashed lines) are shown. The green lines represent loops connecting structural elements (β19-α14, residues 414 to 416; β21-β22, residues 464 to 468). (b) Consensus sequences for the recognition of meso -A 2 pm and l -lysine in various bacteria. The amino-acid numbering is that of Escherichia coli for meso -A 2 pm bacteria and Staphylococcus aureus for l -lysine bacteria. The two conserved tetrapeptide motifs are shown in bold type. According to Dementin (2001) , Gordon (2001) , and Boniface (2006) .

The residues in positions 4 and 5 of the peptide stem are added as a dipeptide by MurF. d -Ala- d -Ala is the most usual dipeptide. d -Ala- d -Ser and d -Ala- d -Lac are found in vancomycin-resistant strains ( Healy et al. , 2000a ). The enzyme from Escherichia coli has been purified and studied ( Duncan et al. , 1990 ; Anderson et al. , 1996 ), and as for MurC, it follows a sequential, ordered kinetic mechanism. The two forms of UDP-MurNAc-tripeptide ( meso -A 2 pm and l -Lys) are equally effective as substrates, and a strong inhibition by excess of UDP-MurNAc-tripeptide has been seen; this effect was suppressed by the addition of 0.5 M NaCl ( Anderson et al. , 1996 ). The specificity profile for the dipeptide substrate has been the subject of many studies; however, because most of them used in vivo systems [see references in van Heijenoort (2001) ], the results obtained are indirect. When the pure enzyme has been available, it was firmly established that it has a high degree of specificity for the C-terminal amino acid ( Duncan et al. , 1990 ; Bugg et al. , 1991 ). This is complementary to the specificity of d -Ala: d -Ala ligase (Ddl), which resides mainly on the N-terminal amino acid, and this constitutes a ‘double sieving’ mechanism that ensures the synthesis of UDP-MurNAc-pentapeptide ending mainly in d -Ala- d -Ala ( Neuhaus & Struve, 1965 ; Duncan et al. , 1990 ). An interesting observation is the ability of MurF to incorporate the dipeptide 3-fluoro- d -Ala-3-fluoro- d -Ala that is synthesized by Ddl in vivo , thereby explaining the autoantagonistic effect of high concentrations of the antibiotic 3-fluoro- d -Ala ( Kollonitsch et al. , 1973 ; Duncan et al. , 1990 ). The MurF enzyme from Thermotoga maritima has been isolated; not unexpectedly, it adds d -Ala- d -Ala to the l -Lys-containing UDP-MurNAc-tripeptide, but not to the d -Lys-containing nucleotide. However, the fact that the latter is a good substrate for MraY explains the incorporation of d -Lys into Thermotoga maritima peptidoglycan ( Boniface et al. , 2006 ).

The crystal structure of the MurF apoenzyme from Escherichia coli has been determined; it is an open structure that is expected to undergo domain closure upon substrate binding ( Yan et al. , 2000 ). Recently, three structures of the enzyme from Streptococcus pneumoniae were cocrystallized with sulfonamide inhibitors ( Fig. 13d ) and showed interdomain closure ( Longenecker et al. , 2005 ; Stamper et al. , 2006 ).

Murein peptide ligase (Mpl) is a nonessential enzyme that is found in some Gram-negative species. The mpl gene was identified by a search of databases for proteins with significant homology with MurC. Mpl participates in the recycling of peptidoglycan by adding the tripeptide l -Ala-γ- d -Glu- meso -A 2 pm onto UDP-MurNAc ( Mengin-Lecreulx et al. , 1996 ). The enzyme from Escherichia coli has been purified recently and its in vitro substrate specificity has been studied thoroughly ( Hervé, 2007 ). The meso -A 2 pm-containing tri-, tetra- and pentapeptides were accepted as substrates with high, similar catalytic efficiencies. Their l -Lys-containing counterparts were accepted with a lower ( c . 500-fold), albeit still significant, efficiency. Weak additions of l -Ala (a MurC-type activity) and l -Ala- d -Glu were seen.

Although not essential, the Mpl enzyme may be interesting as a potential target for antibacterial compounds. Indeed, its broad specificity raises the possibility of incorporating toxic peptides into peptidoglycan. In this regard, Escherichia coli cells were grown in the presence of large concentrations of the synthetic tripeptide l -Ala-γ- d -Glu- l -Lys; however, no effects on cell growth or morphology were seen ( Hervé, 2007 ). This disappointing result was explained by the poor uptake of the lysine-containing peptides by Escherichia coli ( Le Roux, 1991 ). Nevertheless, further work is in progress to identify more permeant Mpl peptide substrates that are endowed with antibacterial activity.

Synthesis of UDP-MurNAc and the UDP-MurNAc-peptides

The UDP-MurNAc and the UDP-MurNAc-peptides were initially isolated from Staphylococcus aureus by Park (1952) . Over the next four decades, these compounds were mainly prepared from bacterial extracts [see references in Flouret (1981) ]. These procedures were tedious and time-consuming; moreover, for some nucleotides (e.g. UDP-MurNAc- l -Ala), the yields were low ( Michaud et al. , 1987 ). Two advances contributed to major improvements in this field: (1) the development of methods of synthesis of UDP-MurNAc and (2) the availability of pure Mur ligases in large quantities.

The first chemical synthesis of UDP-MurNAc was published 40 years ago, although the description was incomplete and the authors could not separate the α and β anomers of the product ( Heymann et al. , 1968 ). Blanot and coworkers described a synthesis starting from commercially available benzyl N -acetyl-4,6- O -benzylidene muramic acid and separated the two anomers of the product by HPLC ( Blanot et al. , 1994 ). This synthesis was improved and scaled up by Dini (2000) . Recently, the synthesis was further modified and optimized using different protecting and coupling strategies ( Babič & Pečar, 2007 ; Kurosu et al. , 2007 ). The key steps of all these synthetic schemes are the introduction of the phosphate group at the anomeric centre of a suitably protected MurNAc derivative, and the coupling of MurNAc-1-phosphate with uridine 5′-phosphomorpholidate.

The in vitro enzymatic synthesis of UDP-MurNAc from UDP-GlcNAc using MurA and MurB was described for the first time by Benson (1993) . The sensitivity of MurB to substrate and product inhibition and the intrinsic NADPH oxidase activity of MurB complicate this procedure. Improvements have been described, though, such as operating under an argon atmosphere ( Reddy et al. , 1999 ) or using an in situ NADPH regeneration system ( Liu et al. , 2001 ). An enzymatic synthesis of UDP-[ 14 C]MurNAc has been published ( Bouhss et al. , 2002 ), while using MurA alone, UDP-GlcNAc- enol pyruvate, the substrate of MurB, can be prepared (Benson et al. , 1993b).

The UDP-MurNAc-peptides are synthesized from UDP-MurNAc by the use of the Mur ligases that act either individually ( Jin et al. , 1996 ; Bertrand et al. , 1997 ; Auger et al. , 1998 ; Raymond et al. , 2003 ) or at the same time ( Reddy et al. , 1999 ; Bouhss et al. , 2004 ; Kurosu et al. , 2007 ). The enzymes from Escherichia coli are used generally, but recently those from Staphylococcus aureus ( Girardin et al. , 2003 ), Pseudomonas aeruginosa ( Paradis-Bleau et al. , 2006 ) and Thermotoga maritima ( Boniface et al. , 2006 ; Babič, 2007 ) have been used. The nonabsolute substrate specificity of the MurE enzymes has been exploited for the synthesis of nucleotides containing unusual amino acids at position 3, such as ll -A 2 pm ( Mengin-Lecreulx et al. , 1988 ), meso -lanthionine and l - allo -cystathionine ( Mengin-Lecreulx et al. , 1994 ), l -ornithine, l -aminopimelate and amidated meso -A 2 pm ( Girardin et al. , 2003 ). Similarly, Schouten (2006) used MurF to prepare UDP-MurNAc-pentapeptide analogues containing d -Cys at position 4 or 5; these derivatives were labelled with pyrene maleimide, thereby yielding fluorescent nucleotide precursors. A total chemical synthesis of UDP-MurNAc-pentapeptide ( Hitchcock et al. , 1998 ; Narayan & VanNieuwenhze, 2007 ), as well as a chemo-enzymatic synthesis of its depsipeptide ( d -Ala- d -Lac-containing) analogue, have been reported ( Liu et al. , 2001 ). Nucleotides modified on the uracil ( Bertrand et al. , 1997 ) and MurNAc ( Ueda et al. , 2004 ) moieties have also been prepared, while radioactive forms of the UDP-MurNAc-peptides have been synthesized from radiolabelled amino acids ( Mengin-Lecreulx et al. , 1988 ; Michaud et al. , 1990 ; Pratviel-Sosa et al. , 1994 ; Bouhss et al. , 2004 ; Boniface et al. , 2006 ).

The analysis and purification of the UDP-MurNAc-peptides have been aided considerably by the introduction of reverse-phase HPLC ( Flouret et al. , 1981 ). As well as for synthesis, this technique has found applications in the measurement of the enzymatic activity of the Mur ligases ( Liger et al. , 1991 ; Auger et al. , 1995 ) and in the analysis of the intracellular pools of these nucleotide precursors ( Mengin-Lecreulx et al. , 1982 ).

For the setting up of in vitro inhibitory assays, studies have used the in situ production of UDP-MurNAc-peptides from UDP-GlcNAc by MurA, MurB and the Mur ligases ( Wong et al. , 1998 ; El Zoeiby, 2001 ). Similarly, a MurF inhibitory assay in which the meso -A 2 pm-containing or l -Lys-containing UDP-MurNAc-tripeptide is produced by Mpl in situ has been described ( Baum et al. , 2006 ). In both cases, HPLC was used to analyse the nucleotides formed.

Inhibitors of the Mur ligases

Several simple compounds that are structurally related to l -Ala have been reported to be moderate inhibitors of MurC from Escherichia coli ( Liger et al. , 1991 , 1995 ). A series of phosphinate transition-state analogues of MurC were prepared and compound 7 ( Fig. 15 ) was identified as its most potent inhibitor, with an IC 50 value of 49 nM. Biochemical characterization revealed that it has a mixed-type inhibition with respect to all three substrates. Any structural modification of this inhibitor significantly reduced the inhibitory activity ( Marmor et al. , 2001 ; Reck et al. , 2001 ). Benzylidene rhodanines inhibited MurC with micromolar IC 50 values (12–27 μM): in a whole-cell assay, they were active against methicillin-resistant Staphylococcus aureus (MRSA), but not against Escherichia coli ( Sim et al. , 2002 ). 2-Phenyl-5,6-dihydro-2 H -thieno[3,2- c ]pyrazol-3-ol derivatives with the general formula 8 showed good inhibitory activities against Staphylococcus aureus MurB (the best IC 50 values ranged from 3.6 to 24 μg mL −1 ), MurC (IC 50 values between 7 and 25 μg mL −1 ) and MurD (IC 50 values between 8.3 and 25 μg mL −1 ) enzymes and promising antimicrobial activities against Gram-positive bacteria, including resistant strains. Also here, the MIC values increased to above 128 μg mL −1 when these compounds were tested in the presence of BSA ( Li et al. , 2003 ). A benzofuran acyl-sulfonamide derivative was discovered by AstraZeneca and shown to act competitively with ATP and UDP-MurNAc (IC 50 =2.3 μM), but unfortunately it also has high-affinity binding to BSA ( Ehmann et al. , 2004 ). More than 180 pulvinones [e.g. 9 : IC 50 (MurC)=8 μg mL −1 )] were synthesized and evaluated as inhibitors of MurA-D. They consistently inhibited MurC (IC 50 values in the 1–10 μg mL −1 range), and also MurA and MurB to lesser extents, while demonstrating antibacterial activities against Gram-positive bacteria, including resistant strains ( Antane et al. , 2006 ). Two peptide inhibitors of MurC from Pseudomonas aeruginosa , with IC 50 values of 1.5 and 0.9 mM, were selected by phage display ( El Zoeiby, 2003b ).

Formulae of inhibitors of the Mur ligases.

The effects of various analogues of d -Glu on MurD from Escherichia coli have been studied, and have yielded some moderate inhibitors ( Pratviel-Sosa et al. , 1994 ). Many phosphonic acids and phosphinates (e.g. 10 : IC 50 =20 nM) have been developed so far as substrate analogues and tetrahedral transition-state analogue inhibitors of MurD, respectively ( Auger et al. , 1995 ; Tanner et al. , 1996 ; Gegnas et al. , 1998 ; Snyder et al. , 1999 ; Victor et al. , 1999 ; Gobec et al. , 2001 ; Štrancar, 2006 ), and quantitative structure–activity relationship (QSAR) studies have been performed for some of these ( Kotnik et al. , 2004 ). From a library of N -acyl- d -Glu derivatives, several inhibitors have been identified; those containing an indole moiety appeared to be of special interest ( Victor et al. , 1999 ). Recently, the high-resolution crystal structures of MurD in complexes with N -sulfonyl- d -Glu (IC 50 =280 μM) and N -sulfonyl- l -Glu (IC 50 =710 μM) ( 11 ) have been solved ( Fig. 16 ). The binding modes of these inhibitors have also been characterized kinetically ( Kotnik et al. , 2007b ). Using a de novo structure-based molecular design, a series of macrocyclic inhibitors 12 were developed, which showed IC 50 values between 0.7 and 5.1 μM ( Horton et al. , 2003 ). In addition, peptide inhibitors of MurD from Pseudomonas aeruginosa (best IC 50 value: 4 μM) have been obtained by the screening of phage display libraries using competitive biopanning approaches ( Paradis-Bleau et al. , 2006 ).

Binding mode of an N -sulfonyl- d -Glu derivative in the active site of Escherichia coli MurD (PDB entry: 1jff). The inhibitor (compound 11 , D isomer, in Fig. 15 ) is shown in magenta. Hydrogen bonds are indicated by dashed lines. The green lines represent sequence elements not directly involved in the interaction.

Analogues of A 2 pm (90% inhibition at 5 mM compound) and some N -acyl-dipeptide derivatives (IC 50 =0.6–10 mM) have shown moderate inhibitory activities on MurE from Escherichia coli ( Abo-Ghalia et al. , 1985 , 1988 ; Michaud et al. , 1990 ; van Assche, 1991 ; Le Roux, 1992 ; Auger et al. , 1996 ). Phosphinates designed as transition-state analogues of MurE inhibited the Escherichia coli enzyme in the micromolar range; the best of these ( 13 ) had an IC 50 value of 1.1 μM, whereas the derivative devoid of the UMP part was a poor inhibitor (IC 50 =700 μM) ( Zeng et al. , 1998 ). In addition, some phosphinates and β-sulfonamidopeptides designed as transition-state analogue inhibitors of MurD can inhibit MurE from Staphylococcus aureus , presumably by acting as substrate analogues ( Humljan et al. , 2006 ; Štrancar, 2007 ).

The first inhibitors of MurF were pseudo-tripeptide and pseudo-tetrapeptide aminoalkylphosphinic acids of the general structure X-Lys-Ψ(PO 2 H)-Gly-Ala that had been synthesized as transition-state analogues. They acted as reversible competitive inhibitors, with K i values in the range of 200–700 μM ( Miller et al. , 1998 ). With the affinity selection screening, two small-molecule MurF leads (e.g. 14 : IC 50 =1 μM) were discovered by Abbott Laboratories and cocrystallized with the enzyme ( Gu et al. , 2004 ; Longenecker et al. , 2005 ; Comess et al. , 2006 ) ( Fig. 13d ). Both these were bound in the substrate-binding region and induced domain closure. After the structure-based lead optimization, a series of potent inhibitors were obtained, culminating in compound 15 (IC 50 =22 nM), although none of these exhibited significant antibacterial activities even in the presence of bacterial cell permeabilizers ( Gu et al. , 2004 ; Stamper et al. , 2006 ). Recently, three QSAR models were constructed to further facilitate the search for new inhibitors with extensive physicochemical properties ( Kong et al. , 2007 ). Using an Mpl-based assay, a thiazolylaminopyrimidine series of MurF inhibitors with IC 50 values as low as 2.5 μM have also been identified ( Baum et al. , 2006 ).

Besides the hexoses and nucleotides described in the previous sections, the cytoplasmic steps of peptidoglycan biosynthesis involve a certain number of substrates ( Fig. 1 ). Most participate in many metabolic pathways (acetyl-CoA, UTP, PEP, NADPH, ATP and l -amino acids), but some are more specific ( meso -A 2 pm, d -Ala, d -Ala- d -Ala and d -Glu). Because meso -A 2 pm is the last intermediate in the l -Lys biosynthesis pathway, which has been the subject of several reviews ( Patte et al. , 1983 ; Cox et al. , 1996 ; Born & Blanchard, 1999 ; Cox et al. , 2000 ; Hutton et al. , 2007 ), it will not be considered here. Therefore, only the formation of d -Ala- d -Ala (and its precursor d -Ala) and d -Glu will be described.

Formation of d -Ala- d- Ala

Essentially found in peptidoglycan, d -Ala is also present in lipoteichoic acids of Gram-positive organisms ( Volkman et al. , 2001 ). It is produced from l -Ala through the action of a pyridoxal 5′-phosphate (PLP)-dependent enzyme, alanine racemase. Some organisms contain only one alanine racemase, while others have two, encoded by the alr and dadX genes. The latter group includes Escherichia coli ( Wild et al. , 1985 ), Salmonella typhimurium ( Wasserman et al. , 1984 ; Esaki & Walsh, 1986 ) and Pseudomonas aeruginosa ( Strych et al. , 2000 ). Expression of the alr gene is constitutive and provides the d -Ala that is necessary to maintain cell growth, while the dadX -encoded racemase is inducible and required only when l -Ala is used as a carbon and energy source. While Alr from Enterococcus faecalis ( Badet & Walsh, 1985 ), Salmonella typhimurium ( Esaki & Walsh, 1986 ), Thermus thermophilus ( Seow et al. , 2000 ), Shigella sp. ( Yokoigawa et al. , 2001 ) and Helicobacter pylori ( Saito et al. , 2007 ) have been described as monomers, those from Escherichia coli , Pseudomonas aeruginosa ( Strych & Benedik, 2002 ), Bifidobacterium bifidum ( Yamashita et al. , 2003 ), Corynebacterium glutamicum ( Oikawa et al. , 2006 ), Acidophilium organovorum ( Seow et al. , 1998 ), Mycobacterium sp. ( Strych et al. , 2001 ) and Bacillus stearothermophilus ( Inagaki et al. , 1986 ) are dimers that are formed between identical polypeptide chains of c . 40 kDa. Structural studies on Alr from Bacillus stearothermophilus ( Shaw et al. , 1997 ; Morollo et al. , 1999 ) and Mycobacterium tuberculosis ( LeMagueres et al. , 2005 ) have established that each monomer consists of two different domains: an N-terminal domain made up of an α/β-barrel, and a C-terminal one primarily made up of β-strands. Each subunit contains one PLP molecule ( Fig. 17 ). Kinetic analyses ( Sawada et al. , 1994 ) and X-ray crystallographic studies ( Shaw et al. , 1997 ; Watanabe et al. , 2002 ) have revealed that the Alr reaction proceeds via a two-base mechanism ( Fig. 18 ). First, the PLP bound to the active-site residue Lys39 ( Bacillus stearothermophilus numbering) reacts with l -Ala to form an external Schiff base through transaldimination. The reaction continues with the abstraction of the C α proton of the external aldimine ( l -Ala-PLP adduct) by Tyr265 from the second monomer. In the next step, the planar carbanion intermediate is reprotonated by Lys39 from the opposite side to yield the d -enantiomer. Finally, the internal aldimine between Lys39 and PLP is formed again, displacing d -Ala from its covalent linkage to the cofactor. Conversion of d - into l -enantiomer proceeds in the opposite manner. The importance of the catalytic bases Lys39 and Tyr265 has been revealed by several studies using mutant enzymes ( Watanabe et al. , 1999a , b ). Replacing Tyr265 by Ala in Alr from Bacillus stearothermophilus resulted in a novel aldolase activity ( Seebeck & Hilvert, 2003 ). Another interaction of great importance for the racemization process is the direct hydrogen bond between the unprotonated pyridine nitrogen of PLP and Arg219 ( Shaw et al. , 1997 ; Morollo et al. , 1999 ). In other PLP-dependent enzymes, but not with Alr, the pyridine nitrogen of the cofactor is in the protonated form and acts as an effective electron sink, stabilizing the carbanion intermediate by forming a quinonoid species. The unusual interaction between the positively charged guanidino group of Arg219 and the pyridine nitrogen of PLP in Alr was explored by analysis of the R219K, R219A and R219E mutants ( Sun & Toney, 1999 ). It was thus demonstrated that for efficient catalysis, a positively charged residue is required in this position. Molecular dynamics simulations have revealed that the enhancement of the carbon acidity of the α-amino acid by PLP via unprotonated pyridine is mainly due to solvation effects, in contrast to the intrinsic electron-withdrawing stabilization by the pyridinium ion to form a quinonoid intermediate ( Major & Gao, 2006 ).

Three-dimensional structure of the dimer of alanine racemase from Mycobacterium tuberculosis (PDB entry: 1xfc). Monomers are shown in blue and green. PLP is shown in magenta. The red spheres represent the side-chain of Lys42 (equivalent to Lys39 of Alr from Bacillus stearothermophilus ).

Two-base mechanism of the reaction catalysed by alanine racemase. Lys39 and Tyr265′ ( Bacillus stearothermophilus numbering) act as the catalytic bases abstracting the α-hydrogen from d -Ala and l -Ala, respectively. The reaction proceeds through a planar carbanionic intermediate (top, right and bottom, left). Owing to the participation of the carboxylate of Ala, Lys39 and Tyr265′ remain unionized throughout the catalytic process. According to Watanabe (2002) .

Cycloserine is a naturally occurring suicide substrate of many PLP-dependent enzymes; in this respect, d -cycloserine ( Fig. 19 ) inactivates Alr in a time-dependent manner ( Wang & Walsh, 1978 ). After the initial formation of an external aldimine between cycloserine and PLP, a proton is transferred from the C-2 of the substrate to the C-4′ of the cofactor, resulting in the formation of a ketimine species. This ketimine proceeds through a second prototropic shift, forming a stable isoxazole ( Fig. 19 ). Together with kinetic studies, the crystal structures of l - and d -cycloserine-inactivated Alr ( Fenn et al. , 2003 ) and the Y265F mutant enzyme ( Fenn et al. , 2005 ) from Bacillus stearothermophilus have emphasized the importance of the catalytic bases Tyr265′ and Lys39 for the inactivation process.

Mechanism of inactivation of alanine racemase by d -cycloserine. Lys39 and Tyr265′ ( Bacillus stearothermophilus numbering) catalyse two proton transfers leading successively to the ketimine intermediate and the final isoxazole derivative. PMP, pyridoxamine phosphate. According to Fenn (2003) .

A series of N (2)-substituted derivatives of compound 16 ( Kim et al. , 2003b ) ( Fig. 20 ) and five- and six-membered heterocycles ( Kim et al. , 2003a ) were prepared and evaluated for inhibitory activities against Alr from various bacterial species, as well as in growth inhibition assays. Some of the heterocycles had moderate inhibitory activities against Alr from Escherichia coli , Staphylococcus aureus , Pseudomonas aeruginosa and Mycobacterium tuberculosis . Besides d -cycloserine, several other compounds are irreversible time-dependent mechanism-based inactivators (suicide substrates) of Alr, like O -carbamyl- d -serine ( Wang & Walsh, 1978 ), β-chloro- and β-fluoro- d -alanine ( Wang & Walsh, 1978 ; Badet et al. , 1984 ), β,β - difluoro- ( Wang & Walsh, 1981 ) and β,β,β-trifluoroalanine 17 ( Faraci & Walsh, 1989 ) and 3-halovinylglycines 18 ( Thornberry et al. , 1987 , 1991 ). Other alanine analogues that are inhibitors of Alr have been reviewed by Neuhaus & Hammes (1981) . To improve in vivo antimicrobial activities, several di- and tripeptides containing β - chloro- l -Ala ( Cheung et al. , 1983 , 1986 ; Boisvert et al. , 1986 ; Le Roux, 1991 ) and halovinylglycine ( Patchett et al. , 1988 ) were synthesized as transport systems for the intracellular delivery of potentially bactericidal amino acids. l -Norvalyl- l -chlorovinylglycine showed good activity against Gram-positive organisms, including methicillin-resistant Staphylococcus species ( Patchett et al. , 1988 ). Furthermore, (β-chloro-, (β-β-dichloro- and (β-β-β-trichloro-α-aminoethy) phosphonic acids ( Vo-Quang et al. , 1986a ) and the phosphonic analogue of vinylglycine, dl -(1-amino-2-propenyl)phosphonic acid ( Vo-Quang et al. , 1986b ), have been evaluated against Alr and Ddl from Pseudomonas aeruginosa and Enterococcus faecalis . The monochloro, dichloro and vinylglycine derivatives of Ala-P exhibited strong inhibition of Alr from both species tested, whereas only the Ddl from Enterococcus faecalis was inhibited by these compounds.

Formulae of inhibitors of alanine racemase and d -Ala- d -Ala ligase.

The phosphonic analogue of alanine, l -1-aminoethylphosphonic acid 19 ( l -AlaP) ( Atherton et al. , 1979b ), is a time-dependent irreversible inactivator of Gram-positive ( Bacillus , Staphylococcus and Streptococcus ), but not Gram-negative ( Escherichia coli , Salmonella and Pseudomonas ), racemases ( Badet & Walsh, 1985 ; Badet et al. , 1986 ). It was used as the phosphonodipeptide alaphosphin ( l -Ala- l -AlaP), which can be transported by bacterial cell wall permeases and then hydrolysed to l -Ala and l -AlaP ( Allen et al. , 1978 ; Atherton et al. , 1979a ). The formation of an AlaP-PLP Schiff base linkage was shown by solid-state 15 N-NMR of the [ 15 N]AlaP-racemase complex ( Copié, 1988 ) and the cocrystal structure of l -AlaP with Alr ( Stamper et al. , 1998 ). The latter showed an inappropriate orientation of the external aldimine for efficient Cα proton abstraction and revealed interactions of the phosphonate group with putative catalytic residues, thereby rendering them unavailable for catalysis. Several phosphonodipeptides containing 1-amino-1-methylethanephosphonic ( Zboinska et al. , 1990 ), 1-aminomethylphosphonic ( Atherton et al. , 1982 ) and 1-aminocyclopropanephosphonic acid 20 ( Erion & Walsh, 1987 ) were generally less potent inhibitors of the Alr and Ddl enzymes than phosphonodipeptides based on AlaP.

The replacement of the AlaP phosphonate with a boronic acid group led to the alanine analogue (1-aminoethyl)boronic acid 21 , which is a slow-binding, time-dependent inhibitor of Alr ( Bacillus stearothermophilus ) and Ddl ( Salmonella typhimurium ). The inhibition is most probably due to the formation of a tetrahedral boronate anion that acts as a transition-state analogue ( Duncan et al. , 1989 ). On the other hand, compounds mimicking the PLP-Ala complex were inactive ( Leung et al. , 1985 ).

A structure-based strategy for the identification of novel Bacillus stearothermophilus Alr inhibitors with a dynamic receptor-based pharmacophore model was developed using the LigBuilder programme ( Mustata & Briggs, 2002 ); however, these compounds have not yet been evaluated for their inhibitory activities.

The condensation of two molecules of d -Ala that were formed previously by alanine racemase is catalysed by a specific ATP-dependent D-Ala:D-Ala ligase (Ddl). Early kinetic and specificity studies of the reaction were carried out with the ligase purified from Enterococcus faecalis ( Neuhaus et al. , 1962a ): they provided evidence for two d -Ala-binding sites that have different specificity patterns and Michaelis constants ( Neuhaus, 1962b ). The binding site for the N-terminal d -Ala, designated as the donor site, is highly specific for d -Ala whereas the C-terminal d -Ala-binding site, the acceptor site, is less specific and accepts a variety of d -amino acids. Ddl enzymes from Oceanobacillus iheyensis , Synechocystis sp. ( Sato et al. , 2005 ) and Thermotoga maritima ( Sato et al. , 2006 ) were also isolated and characterized for their substrate specificities. Recently, the entire MurC-Ddl fusion protein from Chlamydia trachomatis (see ‘ Biosynthesis of the UDP-MurNAc-peptides ’) was purified; the presence of the MurC domain appeared to be required for Ddl activity ( McCoy & Maurelli, 2005 ). The existence of the two distinct genes ddlA and ddlB in Escherichia coli ( Zawadzke et al. , 1991 ; Al-Bar et al. , 1992 ) and Salmonella typhimurium ( Daub et al. , 1988 ) was demonstrated. Both genes were cloned and overexpressed, and their products have been purified. Despite a difference in size, both enzymes show very similar kinetic characteristics and substrate specificities.

The kinetic mechanism, which has been well studied with the enzyme from Salmonella typhimurium , has been shown to be an ordered ter–ter reaction with ATP as the first substrate to bind, and ADP the last product off. The reaction is reversible; however, as the exchange reaction is not strictly ADP-dependent, this can argue in favour of some randomness in the kinetic mechanism ( Mullins et al. , 1990 ).

The reaction starts with the attack on the first d -Ala by the γ-phosphate of ATP, to give an acylphosphate. This is followed by an attack by the amino group of the second d -Ala to yield a tetrahedral intermediate, which collapses into d -Ala- d -Ala and P i ( Mullins et al. , 1990 ; Healy et al. , 2000b ). In this respect, the mechanism is similar to that of the Mur ligases (see ‘ Biosynthesis of the UDP-MurNAc-peptides ’). Ddl is strongly inhibited by its reaction product d -Ala- d -Ala ( Mullins et al. , 1990 ).

Despite the similarity of the Ddl and Mur ligase reactions, Ddl does not belong to the Mur ligase family. Sequence alignments and crystal structures have revealed that it belongs to the ATP-grasp family, which is composed of highly diverse enzymes that catalyse the ATP-dependent ligation of a carboxyl group to an amino or imino nitrogen, a hydroxyl oxygen or a thiol sulphur ( Galperin & Koonin, 1997 ; Kobayashi & Go, 1997 ) ( Fig. 21 ).

Three-dimensional structure of the Escherichia coli DdlB complexed with ADP and the phosphorylated derivative of phosphinate 22 in Fig. 20 (PDB entry: 2dln). The three domains characteristic of the ATP-grasp fold are shown in green, yellow and blue. Ligands are shown in magenta. Two Mg 2+ ions cocrystallized with DdlB are depicted as black spheres.

The most important inhibitor of Ddl is undoubtedly a structural analogue of d -Ala, d -cycloserine (formula in Fig. 19 ) ( Strominger et al. , 1960 ; Neuhaus & Lynch, 1964 ). A series of phosphinates [e.g. 22 : IC 50 ( Streptococcus faecalis )=35 μM] ( Fig. 20 ), phosphonates and phosphonamidates have been developed as transition-state analogue inhibitors or as analogues of d -alanyl phosphate ( Lacoste et al. , 1979 ; Parsons et al. , 1988 ; Chakravarty et al. , 1989 ; Lacoste et al. , 1991 ; Ellsworth et al. , 1996 ). It was shown that transition-state mimetics can be phosphorylated by Ddl and inhibit the reaction by their tight binding to the enzyme after this phosphorylation ( Duncan & Walsh, 1988 ; McDermott et al. , 1990 ). Although their antibacterial activities are low, they enabled the crystallographic determination of complexes of Escherichia coli DdlB with ADP/phosphorylated phosphinate ( Fan et al. , 1994 ) ( Fig. 21 ) and the Escherichia coli Y216F DdlB mutant with ADP/phosphorylated phosphonate ( Fan et al. , 1997 ). Recently, two patent applications have been filed describing structure-based drug discovery methods for identifying inhibitors that target the ATP-binding site as well as new heterocyclic inhibitors ( Moe et al. , 2003 ; Navia et al. , 2003 ). Using a de novo structure-based molecular design, cyclopropane derivative 23 [ K i ( Escherichia coli )=12.5 μM] was developed as an inhibitor of DdlB ( Besong et al. , 2005 ). An allosteric inhibitor of Ddl from Staphylococcus aureus ( 24 : K i =4 μM) was discovered by high-throughput screening, and it was cocrystallized with the enzyme ( Liu et al. , 2006 ). Recently, diazenedicarboxamides [e.g. 25 : IC 50 ( Escherichia coli )=15 μM] were described as inhibitors of DdlB from Escherichia coli ( Kovač, 2007 ). In addition, the crystal structure of the apo form of Ddl from Thermus caldophilus has been solved, providing an insight into the substrate-induced conformational changes, which could be important for inhibitor design ( Lee et al. , 2006a ).

In vancomycin-resistant strains, the d -Ala- d- Ala termini are replaced by either d -Ala- d -Lac or by d -Ala- d -Ser. The affinities of these alternate termini for the antibiotic vancomycin are three and one orders of magnitude, respectively, lower than that of the conventional d -Ala- d -Ala termini. Depsipeptide d -Ala- d -Lac and dipeptide d -Ala- d -Ser are synthesized by enzymes known as VanA-G according to the vancomycin-resistance phenotype considered. Together with Ddl, the Van enzymes constitute a superfamily of d -Ala: d -X ligases (X= d -Ala, d -Lac or d -Ser) that share a common reaction mechanism and the ATP-grasp three-dimensional structure. Differences in the specificity of the acceptor site have been explained through their pH profiles and active-site residues [see Healy (2000a) for a review]. The crystal structures of d -Ala: d -Lac ligases from Leuconostoc mesenteroides and Enterococcus faecium are also available ( Kuzin et al. , 2000 ; Roper et al. , 2000 ).

Formation of d -glutamic acid

In addition to its occurrence in peptidoglycan ( Vollmer et al. , 2008 ), d -glutamic acid is a key component of capsular poly-γ-glutamate of some bacterial genera ( Ashiuchi & Misono, 2002 ; Candela & Fouet, 2006 ). Two distinct enzymes have been identified for the formation of d -glutamate: glutamate racemase (MurI) and d -amino acid aminotransferase ( d -AAT).

Glutamate racemase catalyses the interconversion of d - and l -enantiomers of glutamate ( Doublet et al. , 1993 ). The murI genes (also called racE ) from various bacteria have been cloned. Contrary to the situation in most organisms (including Escherichia coli ) that contain only one glutamate racemase gene [see references in Dodd (2007) ], the Bacillus subtilis and Bacillus anthracis genomes have two genes that have been named racE1 and racE2 for the former species, and racE (also called glr ) and yrpC for the latter ( Ashiuchi et al. , 1999 ; Kimura et al. , 2004 ; Shatalin & Neyfakh, 2005 ).

The MurI racemases from c . 10 species, including Bacillus pumilus , Pediococcus pentosaceus , Lactobacillus fermenti and Escherichia coli , have been purified [see references in Dodd (2007) ], as well as the RacE1 and RacE2 enzymes from Bacillus anthracis ( Dodd et al. , 2007 ; May et al. , 2007 ) and the RacE and YrpC enzymes from Bacillus subtilis ( Ashiuchi et al. , 1998 , 1999 ). Thorough biochemical studies have been performed for some of them. The Escherichia coli MurI racemase requires UDP-MurNAc- l -Ala for its activation, and it has been shown that this nucleotide precursor exerts its effects by increasing both the substrate-binding affinity and the turnover rate of the enzyme ( Doublet et al. , 1994 ; Ho et al. , 1995 ). Therefore, the formation of d -glutamic acid is regulated by UDP-MurNAc- l -Ala and thus adjusted to the requirement of peptidoglycan synthesis. Moreover, an excessive racemization of the large intracellular pool of l -glutamic acid is avoided ( Doublet et al. , 1993 , 1994 ). This activating mechanism appears to be unique for Escherichia coli because glutamate racemases from other bacteria are not activated by UDP-MurNAc- l -Ala ( Yoshimura & Esaki, 2003 ; Lundqvist et al. , 2007 ). It was hypothesized that a 21-amino-acid extension at the N-terminus of the Escherichia coli enzyme, which is the major difference between Escherichia coli and other bacteria, was responsible for the nucleotide-mediated activation ( Doublet et al. , 1994 ; Ho et al. , 1995 ); however, the N-terminal-truncated Escherichia coli enzyme retained the activation, although to lesser extents ( Ho et al. , 1995 ; Doublet et al. , 1996 ). The glutamate racemase from Pediococcus pentosaceus is inhibited by haemin due to the formation of a stoichiometric complex ( Choi et al. , 1994 ). Recently, it was shown that glutamate racemases from Escherichia coli , Bacillus subtilis (YrpC, but not RacE) and Mycobacterium tuberculosis inhibit DNA gyrase activity; in the case of Escherichia coli , the presence of the nucleotide precursor was required ( Ashiuchi et al. , 2002 , 2003 ; Sengupta et al. , 2006 ).

Glutamate racemase belongs to a small family of amino acid racemases and epimerases ( Tanner, 2002 ; Yoshimura & Esaki, 2003 ). While most amino acid racemases use PLP for abstraction of the α-proton, glutamate racemase operates without any cofactor or metal ion ( Gallo et al. , 1993 ; Glavas & Tanner, 2001 ). Instead, a ‘two-base’ mechanism that involves two cysteine residues has been suggested ( Fig. 22 ). A thiolate from one of the Cys residues promotes the deprotonation of the substrate at the Cα position, to yield a carbanionic intermediate; then reprotonation by the other Cys residue occurs in the opposite stereochemical sense ( Gallo et al. , 1993 ; Glavas & Tanner, 1999 , 2001 ). The catalytic mechanism of glutamate racemases from Aquifex pyrophilus , Lactobacillus fermenti and Bacillus subtilis has been investigated closely ( Hwang et al. , 1999 ; Glavas & Tanner, 2001 ; Ruzheinikov et al. , 2005 ). Two possibilities, which differ in the protonation states of the catalytic bases prior to the initial deprotonation, have been proposed ( Glavas & Tanner, 2001 ; Ruzheinikov et al. , 2005 ). Furthermore, stereochemistry, substrate ligation and active-site protonation states have been investigated by molecular dynamics simulations ( Möbitz & Bruice, 2004 ; Puig et al. , 2005 ). Recently, the results from computational simulations on a quantum mechanical/molecular mechanical potential energy surface ( Puig et al. , 2006 ) indicated two possible roles for MurI as a catalyst and supported the mechanistic proposal by Rios (2000) for the PLP-independent amino acid racemases.

Reaction mechanism of the MurI enzyme. The enzyme employs two active-site cysteine residues as acid/base catalysts. A thiolate from one of the cysteines abstracts the α-proton, forming a carbanionic intermediate (central formula). The other cysteine thiol delivers a proton to the opposite face of the intermediate. According to Gallo (1993) .

Several crystal structures of bacterial glutamate racemases have been reported ( Hwang et al. , 1999 ; Ruzheinikov et al. , 2005 ; Kim et al. , 2007 ; Lundqvist et al. , 2007 ; May et al. , 2007 ) ( Fig. 23 ). Along with analytical methods such as size-exclusion chromatography or equilibrium ultracentrifugation, structure examination has led to a classification of glutamate racemases according to their quaternary structure ( Lundqvist et al. , 2007 ; May et al. , 2007 ): monomer ( Escherichia coli , Lactobacillus fermenti , Bacillus subtilis YrpC) ( Fig. 23a ), monomer–dimer equilibrium ( Aquifex pyrophilus , Bacillus anthracis RacE1 and Bacillus subtilis RacE) or dimer ( Helicobacter pylori , Enterococcus faecalis , Enterococcus faecium , Staphylococcus aureus and Bacillus anthracis RacE2). Among dimers, a tail-to-tail orientation (i.e. with the active sites opposed and fully exposed to the solvent) is the general case; however, the Helicobacter pylori enzyme appears as a head-to-head dimer (i.e. with the active sites in close proximity in a face-to-face orientation) ( Fig. 23b ).

Three-dimensional structures of two glutamate racemases. (a) Escherichia coli MurI (PDB entry: 2jfn) The protein appears as a monomer. Substrate d -Glu and activator UDP-MurNAc- l -Ala are shown in magenta. (b) Helicobacter pylori MurI (PDB entry: 2jfx). The protein appears as a head-to-head dimer. The two monomers are shown in green and blue. Substrate d -Glu is shown in magenta. Glu151 and Lys117′ side-chains are shown in red; the arrow indicates a putative intermonomer salt bridge contact between them.

Owing to the essential role of glutamate racemase for the viability of bacteria, this enzyme is an attractive target for the development of antibacterial agents. Several inhibitors of glutamate racemase have been reported to date ( Fig. 24 ). Ashiuchi (1993) showed that l -serine O -sulphate 26 behaves as a suicide substrate of the enzyme from Pediococcus pentosaceus ; a mechanism by which an α-aminoacrylate intermediate reacts with a nucleophilic group of the enzyme was proposed. Tanner and coworkers synthesized aziridino glutamate 27 , which irreversibly inactivates Lactobacillus fermenti glutamate racemase through the alkylation of a Cys residue in the active site ( Tanner & Miao, 1994 ), and d - N -hydroxyglutamate 28 , which was shown to be a competitive inhibitor ( K i =56 μM) ( Glavas & Tanner, 1997 ). The research group at Eli Lilly discovered the first potent competitive inhibitors that showed antibacterial activity: 4-substituted d -Glu analogues with various aryl and heteroaryl substituents (e.g. 29 ), with IC 50 values as low as 30 nM and a good correlation between inhibitory and antibacterial activities against Streptococcus pneumoniae ( de Dios, 2002 ). Pyrazolopyrimidinedione derivative 30 was discovered from a high-throughput screen by AstraZeneca; it behaved as an uncompetitive inhibitor of Helicobacter pylor MurI (IC 50 =1.4 μM) and prevented the growth of Helicobacter pylori cells (MIC=4 μg mL −1 ) ( Lundqvist et al. , 2007 ). Kim (2000) isolated peptide ligands from a random phage display library that inhibit Escherichia coli glutamate racemase activity. Among these, the peptide sequence His-Pro-Trp-His-Lys-Lys-His-Pro-Asp-Arg-Lys-Thr has the highest affinity for the enzyme (IC 50 =160 μM); however, no bactericidal or antibacterial effects were seen.

Formulae of inhibitors of MurI.

d -Amino acid aminotransferase ( d -AAT) yields d -glutamate and pyruvate from d -alanine and α-ketoglutarate ( Fig. 25 ) ( Fotheringham et al. , 1998 ), and d -AAT activity has been reported for various Gram-positive bacteria: Bacillus ( Tanizawa et al. , 1989 ; Taylor & Fortheringham, 1997 ; Fotheringham et al. , 1998 ; Peisach et al. , 1998 ; Berger et al. , 2003 ), Staphylococcus ( Pucci et al. , 1995 ) and Listeria ( Thompson et al. , 1998 ). Recently, the d -AATs from Geobacillus sp. were cloned and characterized for their genetic, catalytic and structural aspects ( Lee et al. , 2006b ). The three-dimensional structures of various forms of d -AAT from Bacillus sp. YM-1 have been solved ( Sugio et al. , 1995 , 1998 ; Peisach et al. , 1998 ; van Ophem, 1999 ) and the pathway of the enzymatic reaction has been described in structural terms. Tyr30, Arg98 and His100 are the major residues relating to the stereospecificity of d -AAT, whereby it specifically carries out a transamination reaction using PLP as a cofactor and a catalytic lysine as a general base/acid. The accepted pathway for transamination, which has been characterized for a number PLP-dependent enzymes, proceeds through a ping-pong kinetic mechanism.

Equation of the d -AAT reaction (top) and formulae of d -AAT inhibitors (bottom).

A number of compounds have been reported to be inhibitors of d -AAT activity ( Neuhaus & Hammes, 1981 ; Axelsson et al. , 1994 ). β-Chloro- d -alanine 31 and d -serine- O -sulphate 32 ( Fig. 25 ) are suicide substrates; they are converted into a putative aminoacrylate intermediate, which either reacts with PLP to form a stable adduct or is protonated and hydrolysed to yield pyruvate ( Axelsson et al. , 1994 ; Adams et al. , 2005 ).

Both the transaminase and the glutamate racemase routes have been reported for some bacterial species: Staphylococcus haemolyticus ( Pucci et al. , 1995 ), Bacillus sphaericus ( Fotheringham et al. , 1998 ) and Bacillus subtilis ( Wipat et al. , 1996 ). In other organisms, such as Escherichia coli, Lactobacillus sp. and Pediococcus sp., only the glutamate racemase activity exists ( Nakajima et al. , 1986 ; Doublet et al. , 1993 ; Gallo & Knowles, 1993 ).

The organization of the genes involved in the biosynthetic process reviewed here is well established in Escherichia coli , where it has been facilitated by the isolation of thermosensitive mutants. Most of the genes are clustered in two regions, known as mra and mrb (murein regions A and B, respectively) ( Miyakawa et al. , 1972 ). The mra (or dcw , division cell wall) region is located at 2 min on the chromosome and contains genes involved either in peptidoglycan synthesis or in cell division in the order: mraZ-mraW-ftsL-ftsI-murE-murF-mraY-murD-ftsW-murG-murC-ddlB-ftsQ-ftsZ-envA ( Mengin-Lecreulx et al. , 1989 ). The first nine genes are under the control of the P mra promoter ( Mengin-Lecreulx et al. , 1998 ). The mrb region is located at 90 min and contains the murI and murB genes. The glmM and murA genes are in the 69 min region, and the glmU and glmS genes in the 83 min region. The ddlA , dadX , alr and mpl genes are located at 8.5, 27, 92 and 96 min, respectively ( Berlyn et al. , 1996 ). The mra region has been located in other bacteria such as Bacillus subtilis , Staphylococcus aureus , Enterococcus faecalis and Pseudomonas aeruginosa ( Daniel & Errington, 1993 ; Pucci et al. , 1997 ; El Zoeiby, 2000 ; Azzolina et al. , 2001 ); in the last of these, its organization is exactly the same as in Escherichia coli .

Peptidoglycan integrity being necessary for the bacterial cell's survival, genes involved in its biosynthesis must be essential provided no isogenes are present. Data about the essential character of the genes coding for the cytoplasmic enzymes can be found either in former literature [see references in van Heijenoort (2001) ], or inferred from the recent systematic construction of single-gene knock-out mutants of Escherichia coli ( Baba et al. , 2006 ). In this organism, the glmS , glmM , glmU , murA , murB , murC , murD , murE , murF and murI genes are essential. Isogenes ( ddlA / ddlB , alr / dadX ) can be individually inactivated, showing that the remaining isozyme is sufficient to sustain growth ( Wild et al. , 1985 ; Zawadzke et al. , 1991 ). As other genes involved in the recycling pathway, the mpl gene is dispensable for Escherichia coli growth ( Mengin-Lecreulx et al. , 1996 ).

A systematic gene inactivation was also performed in Bacillus subtilis ( Kobayashi et al. , 2003 ). The glmS , ybbT ( glmM ), gcaD ( glmU ), murA , murB , murC , murD , murE , alr , ddl and racE genes could not be inactivated. As far as alr and ddl are concerned, this is explained because Bacillus subtilis , contrary to Escherichia coli , contains only one alanine racemase and one Ddl. The racE gene was claimed to be essential as no knockout mutants could be found on solid medium, suggesting that yrpC cannot take over. However, the disruption of the racE gene has been described recently ( Ashiuchi et al. , 2007 ); the RacE-less mutants obtained can grow with a reduced rate in liquid medium even in the absence of exogenous d -glutamate. The authors concluded that the RacE activity, although important for ensuring maximum growth rate, is dispensable, and that the YrpC racemase probably operates as an anaplerotic enzyme for RacE ( Ashiuchi et al. , 2007 ).

Although deletion of both murA1 and murA2 genes in Streptococcus pneumoniae is lethal, each can be inactivated individually without effect on cell growth ( Du et al. , 2000 ). In Bacillus anthracis , while the racE1 knock-out leads to moderate growth defect that can be alleviated fully by d -glutamate, the racE2 knock-out severely inhibits bacterial growth, which is only partially restored by exogenous d -glutamate ( Shatalin & Neyfakh, 2005 ).

It is important to mention that some mutations in genes involved in the cytoplasmic steps in MRSA affect their susceptibility to methicillin; this is the case for glmS ( Komatsuzawa et al. , 2004 ), glmM ( Jolly et al. , 1997 ; Glanzmann et al. , 1999 ) and murE ( Ludovice et al. , 1998 ). In Pseudomonas aeruginosa , a transcriptional repressor of the glmS expression, known as glmR , has been described. The inactivation of the glmR gene dramatically sensitizes the microorganism to a large variety of antibiotics, and in particular the aminoglycosides ( Ramos-Aires et al. , 2004 ).

Since the discovery of Park's nucleotide in 1952, the cytoplasmic steps of peptidoglycan biosynthesis have been defined completely. In particular, considerable progress was made during the 1990s, where the overproduction and purification of the enzymes became possible, along with the production of the noncommercially available substrates by chemical and/or enzymatic synthesis. Studies using enzyme kinetics, isotope-based experiments, NMR, sequence alignments, site-directed mutagenesis, etc., have allowed the reaction mechanisms to be established and their substrate specificities to be defined. More recently, the availability of X-ray and NMR structures has improved our knowledge of these mechanisms; furthermore, these investigations have revealed important conformational changes upon ligand binding. Generally, they were performed with a representative enzyme, and often that of Escherichia coli . In the future, it will be important to compare enzymes from different species with respect to ligand binding and reaction mechanism. This is obvious when one of the substrates differs (e.g. MurE from Gram-negative vs. Gram-positive species, d -Ala: d -X ligases); however, even when the substrates are invariant, significant differences may prevail (e.g. MurB types I and II, alanine racemases). Such comparative enzymology of bacterial species needs to be performed.

As peptidoglycan is an essential component of the cell wall of eubacteria, these enzymes represent targets for antibacterial agents. However, they have been underexploited so far, perhaps due to the very few natural antibiotics that are known to inhibit them (bacilysin, fosfomycin, d -cycloserine). The proliferation of resistant bacterial strains renders the search for new antibacterial compounds urgent, and in this respect, these enzymes involved in the cytoplasmic steps need to be considered seriously. Although good inhibitors have been described recently, most are devoid of antibacterial activity due to their inability to cross the cytoplasmic membrane, and thereby to reach their target. The challenge for the next few years will be to design compounds that are endowed with penetration properties while retaining their affinities for their respective targets.

This work was supported by the European Commission through the EUR-INTAFAR project (LSHM-CT-2004-512138), the Centre National de la Recherche Scientifique (UMR 8619 and PICS 3729), the Délégation Générale pour l'Armement (Contrats Jeune Chercheur 036000104 and 056000030 to A.B.), the Ministry of Education, Science and Sports of the Republic of Slovenia, the Franco-Slovene Proteus programme and the Institut Charles Nodier (Ljubljana). The authors thank Samo Turk for help in preparing protein figures, and Chris Berrie for critically reading the manuscript. They apologize to those researchers whose work could not be quoted because of limited space.

Protein figures presented in this review were prepared with PyMol [DeLano WL, The PyMol Molecular Graphics System (2002), DeLano Scientific, Palo Alto, CA, USA; http://www.pymol.org ].

Although its essential character had been demonstrated by Mengin-Lecreulx & van Heijenoort (1996) , glmM (referred to in the ‘Genetic organization’ section above) was annotated as nonessential in the article of Baba (2006) . As a matter of fact, it has been proved that the glmM strain of Baba et al. contains an intact copy of the glmM gene (D. Mengin-Lecreulx, pers. commun.).

Abo-Ghalia M Michaud C Blanot D Van Heijenoort J ( 1985 ) Specificity of the uridine-diphosphate- N -acetylmuramyl-l-alanyl-d-glutamate: meso-2,6-diaminopimelate synthetase from Escherichia coli . Eur J Biochem 153 : 81 – 87 .

Google Scholar

Abo-Ghalia M Flegel M Blanot D Van Heijenoort J ( 1988 ) Synthesis of inhibitors of the meso -diaminopimelate-adding enzyme from Escherichia coli . Int J Peptide Protein Res 32 : 208 – 222 .

Adams B Lowpetch K Thorndycroft F Whyte SM Young DW ( 2005 ) Stereochemistry of reactions of the inhibitor/substrates l- and d-β-chloroalanine with β-mercaptoethanol catalysed by l-aspartate aminotransferase and d-amino acid aminotransferase respectively . Org Biomol Chem 3 : 3357 – 3364 .

Al-Bar OAM O'Connor CD Giles IG Akhtar M ( 1992 ) d-alanine: d-alanine ligase of Escherichia coli . Expression, purification and inhibitory studies on the cloned enzyme . Biochem J 282 : 747 – 752 .

Allen JG Atherton FR Hall MJ Hassall CH Holmes SW Lambert RW Nisbet LJ Ringrose PS ( 1978 ) Phosphonopeptides, a new class of synthetic antibacterial agents . Nature 272 : 56 – 58 .

Anderson MS Eveland SS Onishi HR Pompliano DL ( 1996 ) Kinetic mechanism of the Escherichia coli UDPMurNAc-tripeptide d-alanyl-d-alanine-adding enzyme: use of a glutathione S-transferase fusion . Biochemistry 35 : 16264 – 16269 .

Andres CJ Bronson JJ D'Andrea SV et al. ( 2000 ) 4-Thiazolidinones: novel inhibitors of the bacterial enzyme MurB . Bioorg Med Chem Lett 10 : 715 – 717 .

Antane S Caufield CE Hu W et al. ( 2006 ) Pulvinones as bacterial cell wall biosynthesis inhibitors . Bioorg Med Chem Lett 16 : 176 – 180 .

Anwar RA Vlaovic M ( 1986 ) UDP- N -acetylmuramoyl-l-alanyl-d-glutamyl-l-lysine synthetase from Bacillus sphaericus : activation by potassium phosphate . Biochem Cell Biol 64 : 297 – 303 .

Ashiuchi M Misono H ( 2002 ) Biochemistry and molecular genetics of poly-γ-glutamate synthesis . Appl Microbiol Biotechnol 59 : 9 – 14 .

Ashiuchi M Yoshimura T Esaki N Ueno H Soda K ( 1993 ) Inactivation of glutamate racemase of Pediococcus pentosaceus by l-serine- O -sulfate . Biosci Biotechnol Biochem 57 : 1978 – 1979 .

Ashiuchi M Tani K Soda K Misono H ( 1998 ) Properties of glutamate racemase from Bacillus subtilis IFO 3336 producing poly-γ-glutamate . J Biochem (Tokyo) 123 : 1156 – 1163 .

Ashiuchi M Soda K Misono H ( 1999 ) Characterization of yrpC gene product of Bacillus subtilis IFO 3336 as glutamate racemase isozyme . Biosci Biotechnol Biochem 63 : 792 – 798 .

Ashiuchi M Kuwana E Yamamoto T Komatsu K Soda K Misono H ( 2002 ) Glutamate racemase is an endogenous DNA gyrase inhibitor . J Biol Chem 277 : 39070 – 39073 .

Ashiuchi M Kuwana E Komatsu K Soda K Misono H ( 2003 ) Differences in effects on DNA gyrase activity between two glutamate racemases of Bacillus subtilis , the poly-γ-glutamate synthesis-linking Glr enzyme and the YrpC (MurI) isozyme . FEMS Microbiol Lett 223 : 221 – 225 .

Ashiuchi M Nishikawa Y Matsunaga K Yamamoto M Shimanouchi K Misono H ( 2007 ) Genetic design of conditional d-glutamate auxotrophy for Bacillus subtilis : use of a vector-borne poly-γ-glutamate synthetic system . Biochem Biophys Res Commun 362 : 646 – 650 .

Atherton FR Hall MJ Hassall CH Lambert RW Ringrose PS ( 1979a ) Phosphonopeptides as antibacterial agents: rationale, chemistry, and structure-activity relationships . Antimicrob Agents Chemother 15 : 677 – 683 .

Atherton FR Hall MJ Hassall CH Lambert RW Lloyd WJ Ringrose PS ( 1979b ) Phosphonopeptides as antibacterial agents: mechanism of action of alaphosphin . Antimicrob Agents Chemother 15 : 696 – 705 .

Atherton FR Hall MJ Hassall CH Lambert RW Lloyd WJ Ringrose PS Westmacott D ( 1982 ) Antibacterial activity and mechanism of action of phosphonopeptides based on aminomethylphosphonic acid . Antimicrob Agents Chemother 22 : 571 – 578 .

Auger G Van Heijenoort J Blanot D Deprun C ( 1995 ) Synthesis of N -acetylmuramic acid derivatives as potential inhibitors of the d-glutamic acid-adding enzyme . J Prakt Chem 337 : 351 – 357 .

Auger G Van Heijenoort J Vederas JC Blanot D ( 1996 ) Effect of analogues of diaminopimelic acid on the meso -diaminopimelate-adding enzyme from Escherichia coli . FEBS Lett 391 : 171 – 174 .

Auger G Martin L Bertrand J Ferrari P Fanchon E Vaganay S Pétillot Y Van Heijenoort J Blanot D Dideberg O ( 1998 ) Large-scale preparation, purification, and crystallization of UDP- N -acetylmuramoyl-l-alanine: d-glutamate ligase from Escherichia coli . Protein Expr Purif 13 : 23 – 29 .

Axelsson BS Floss HG Lee SG Saeed A Spencer PA Young DW ( 1994 ) Stereochemistry of conversion of the suicide substrates β-chloro-d-alanine and d- and l-serine O -sulfates into pyruvate by d-amino acid aminotransferase and by l-aspartate aminotransferase . J Chem Soc Perkin Trans 1 2137 – 2142 .

Azzolina BA Yuan X Anderson MS El-Sherbeini M ( 2001 ) The cell wall and cell division gene cluster in the mra operon of Pseudomonas aeruginosa : cloning, production, and purification of active enzymes . Protein Expr Purif 21 : 393 – 400 .

Baba T Ara T Hasegawa M Takai Y Okumura Y Baba M Datsenko KA Tomita M Wanner BL Mori H ( 2006 ) Construction of Escherichia coli K-12 in-frame, single-gene knockout mutants: the Keio collection . Mol Syst Biol 2 : 1 – 11 .

Babič A Pečar S ( 2007 ) An improved total synthesis of UDP- N -acetyl-muramic acid . Tetrahedron Lett 48 : 4403 – 4405 .

Babič A Patin D Boniface A Hervé M Mengin-Lecreulx D Pečar S Gobec S Blanot D ( 2007 ) Chemoenzymatic synthesis of the nucleotide substrates of the Mur ligases. 5th Joint Meeting on Medicinal Chemistry , June 17–21, Portorož, Slovenia, (Kikelj D, ed.), pp. 1–4. Medimond Srl, Bologna, Italy.

Bachelier A Mayer R Klein CD ( 2006 ) Sesquiterpene lactones are potent and irreversible inhibitors of the antibacterial target enzyme MurA . Bioorg Med Chem Lett 16 : 5605 – 5609 .

Badet B Walsh C ( 1985 ) Purification of an alanine racemase from Streptococcus faecalis and analysis of its inactivation by (1-aminoethyl)phosphonic acid enantiomers . Biochemistry 24 : 1333 – 1341 .

Badet B Roise D Walsh CT ( 1984 ) Inactivation of the dadB Salmonella typhimurium alanine racemase by D and L isomers of β-substituted alanines: kinetics, stoichiometry, active site peptide sequencing, and reaction mechanism . Biochemistry 23 : 5188 – 5194 .

Badet B Inagaki K Soda K Walsh CT ( 1986 ) Time-dependent inhibition of Bacillus stearothermophilus alanine racemase by (1-aminoethyl)phosphonate isomers by isomerization to noncovalent slowly dissociating enzyme-(1-aminoethyl)phosphonate complexes . Biochemistry 25 : 3275 – 3282 .

Badet B Vermoote P Haumont PY Lederer F Le Goffic F ( 1987 ) Glucosamine synthetase from Escherichia coli : purification, properties, and glutamine-utilizing site location . Biochemistry 26 : 1940 – 1948 .

Badet B Vermoote P Le Goffic F ( 1988 ) Glucosamine synthetase from Escherichia coli : kinetic mechanism and inhibition by N3-fumaroyl-l-2,3-diaminopropionic derivatives . Biochemistry 27 : 2282 – 2287 .

Badet-Denisot MA René L Badet B ( 1993 ) Mechanistic investigations on glucosamine-6-phosphate synthase . Bull Soc Chim Fr 130 : 249 – 255 .

Badet-Denisot MA Leriche C Massière F Badet B ( 1995 ) Nitrogen transfer in E. coli glucosamine-6-phosphate synthase investigations using substrate and bisubstrate analogs . Bioorg Med Chem Lett 5 : 815 – 820 .

Badet-Denisot MA Fernandez-Herrero LA Berenguer J Ooi T Badet B ( 1997 ) Characterization of l-glutamine: d-fructose-6-phosphate amidotransferase from an extreme thermophile Thermus thermophilus HB8 . Arch Biochem Biophys 337 : 129 – 136 .

Barbosa MDFS Yang G Fang J Kurilla MG Pompliano DL ( 2002 ) Development of a whole-cell assay for peptidoglycan biosynthesis inhibitors . Antimicrob Agents Chemother 46 : 943 – 946 .

Baum EZ Montenegro DA Licata L Turchi I Webb GC Foleno BD Bush K ( 2001 ) Identification and characterization of new inhibitors of the Escherichia coli MurA enzyme . Antimicrob Agents Chemother 45 : 3182 – 3188 .

Baum EZ Crespo-Carbone SM Abbanat D Foleno B Maden A Goldschmidt R Bush K ( 2006 ) Utility of muropeptide ligase for identification of inhibitors of the cell wall biosynthesis enzyme MurF . Antimicrob Agents Chemother 50 : 230 – 236 .

Bearne SL ( 1996 ) Active site-directed inactivation of Escherichia coli glucosamine-6-phosphate synthase. Determination of the fructose 6-phosphate binding constant using a carbohydrate-based inactivator . J Biol Chem 271 : 3052 – 3057 .

Bearne SL Blouin C ( 2000 ) Inhibition of Escherichia coli glucosamine-6-phosphate synthase by reactive intermediate analogues. The role of the 2-amino function in catalysis . J Biol Chem 275 : 135 – 140 .

Benson TE Marquardt JL Marquardt AC Etzkorn FA Walsh CT ( 1993 ) Overexpression, purification, and mechanistic study of UDP- N -acetylenolpyruvylglucosamine reductase . Biochemistry 32 : 2024 – 2030 .

Benson TE Filman DJ Walsh CT Hogle JM ( 1995 ) An enzyme-substrate complex involved in bacterial cell wall biosynthesis . Nat Struct Biol 2 : 644 – 653 .

Benson TE Walsh CT Hogle JM ( 1996 ) The structure of the substrate-free form of MurB, an essential enzyme for the synthesis of bacterial cell walls . Structure 4 : 47 – 54 .

Benson TE Walsh CT Hogle JM ( 1997a ) X-ray crystal structures of the S229A mutant and wild-type MurB in the presence of the substrate enolpyruvyl-UDP- N -acetylglucosamine at 1.8-Å resolution . Biochemistry 36 : 806 – 811 .

Benson TE Walsh CT Massey V ( 1997b ) Kinetic characterization of wild-type and S229A mutant of MurB: evidence for the role of Ser 229 as a general catalyst . Biochemistry 36 : 796 – 805 .

Benson TE Harris MS Choi GH Cialdella JI Herberg JT Martin JP Jr Baldwin ET ( 2001 ) A structural variation for MurB: X-ray crystal structure of Staphylococcus aureus UDP-N-acetylenolpyruvylglucosamine reductase (MurB) . Biochemistry 40 : 2340 – 2350 .

Berger BJ English S Chan G Knodel MH ( 2003 ) Methionine regeneration and aminotransferases in Bacillus subtilis , Bacillus cereus , and Bacillus anthracis . J Bacteriol 185 : 2418 – 2431 .

Berlyn KB Low KB Rudd KE ( 1996 ) Linkage map of Escherichia coli K-12, edition 9 . Escherichia coli and Salmonella: Cellular and Molecular Biology , Vol. 2 ( Neidhardt FC Curtiss R III Ingraham JL Lin ECC Low KB Magasanik B Reznikoff WS Riley M Schaechter M & Umbarger HE , eds), pp. 1715 – 1902 . ASM Press , Washington, DC.

Google Preview

Bertrand JA Auger G Fanchon E Martin L Blanot D Van Heijenoort J Dideberg O ( 1997 ) Crystal structure of UDP- N -acetylmuramoyl-l-alanine: d-glutamate ligase from Escherichia coli . EMBO J 16 : 3416 – 3425 .

Bertrand JA Auger G Martin L Fanchon E Blanot D Le Beller D Van Heijenoort J Dideberg O ( 1999 ) Determination of the MurD mechanism through crystallographic analysis of enzymes complexes . J Mol Biol 289 : 579 – 590 .

Bertrand JA Fanchon E Martin L Chantalat L Auger G Blanot D Van Heijenoort J Dideberg O ( 2001 ) ‘Open’ structures of MurD: domain movements and structural similarities with folylpolyglutamate synthetase . J Mol Biol 301 : 1257 – 1266 .

Besong GE Bostock JM Stubbings W Chopra I Roper DI Lloyd AJ Fishwick CW Johnson AP ( 2005 ) A de novo designed inhibitor of d-Ala-d-Ala ligase from E. coli . Angew Chem Int Ed Engl 44 : 6403 – 6406 .

Blanot D Auger G Liger D Van Heijenoort J ( 1994 ) Synthesis of α and β anomers of UDP- N -acetylmuramic acid . Carbohydr Res 252 : 107 – 115 .

Blewett AM Lloyd AJ Echalier A Fülöp V Dowson CG Bugg TDH Roper DI ( 2004 ) Expression, purification, crystallization and preliminary characterization of uridine 5′-diphospho- N -acetylmuramoyl l-alanyl-d-glutamate: lysine ligase (MurE) from Streptococcus pneumoniae 110K/70 . Acta Cryst D60 : 359 – 361 .

Boisvert W Cheung KS Lerner SA Johnston M ( 1986 ) Mechanisms of action of chloroalanyl antibacterial peptides. Identification of the intracellular enzymes inactivated on treatment of Escherichia coli JSR-O with the dipeptide βCl-lAla-βCl-lAla . J Biol Chem 261 : 7871 – 7878 .

Boniface A ( 2007 ). Etude des relations structure-activité au sein de la famille des Mur synthétases, enzymes de la voie de biosynthèse du peptidoglycane . Ph.D. Thesis, Université Paris-Sud, Orsay, France.

Boniface A Bouhss A Mengin-Lecreulx D Blanot D ( 2006 ) The MurE synthetase from Thermotoga maritima is endowed with an unusual d-lysine adding activity . J Biol Chem 281 : 15680 – 15686 .

Born TL Blanchard JS ( 1999 ) Structure/function studies on enzymes in the diaminopimelate pathway of bacterial cell wall biosynthesis . Curr Opin Chem Biol 3 : 603 – 613 .

Bouhss A Mengin-Lecreulx D Blanot D Van Heijenoort J Parquet C ( 1997 ) Invariant amino acids in the Mur peptide synthetases of bacterial peptidoglycan synthesis and their modification by site-directed mutagenesis in the UDP-MurNAc: l-alanine ligase from Escherichia coli . Biochemistry 36 : 11556 – 11563 .

Bouhss A Dementin S Van Heijenoort J Parquet C Blanot D ( 1999a ) Formation of adenosine 5′-tetraphosphate from the acyl phosphate intermediate: a difference between the MurC and MurD synthetases of Escherichia coli . FEBS Lett 453 : 15 – 19 .

Bouhss A Dementin S Parquet C Mengin-Lecreulx D Bertrand JA Le Beller D Dideberg O Van Heijenoort J Blanot D ( 1999b ) Role of the ortholog and paralog amino acid invariants in the active site of the UDP-MurNAc-l-alanine: d-glutamate ligase (MurD) . Biochemistry 38 : 12240 – 12247 .

Bouhss A Dementin S Van Heijenoort J Parquet C Blanot D ( 2002 ) MurC and MurD synthetases of peptidoglycan biosynthesis: borohydride trapping of acyl-phosphate intermediates . Methods Enzymol 354 : 189 – 196 .

Bouhss A Crouvoisier M Blanot D Mengin-Lecreulx D ( 2004 ) Purification and characterization of the bacterial MraY translocase catalyzing the first membrane step of peptidoglycan biosynthesis . J Biol Chem 279 : 29974 – 29980 .

Bouhss A Trunkfield AE Bugg TDH Mengin-Lecreulx D ( 2008 ) The biosynthesis of peptidoglycan lipid-linked intermediates . FEMS Microbiol Rev , doi: DOI: 10.1111/j.1574-6976.2007.00089.x

Bronson JJ DenBleyker KL Falk PJ Mate RA Ho HT Pucci MJ Snyder LB ( 2003 ) Discovery of the first antibacterial small molecule inhibitors of MurB . Bioorg Med Chem Lett 13 : 873 – 875 .

Broschat KO Gorka C Page JD Martin-Berger CL Davies MS Huang H Gulve EA Salsgiver WJ Kasten TP ( 2002 ) Kinetic characterization of human glutamine-fructose-6-phosphate amidotransferase I. Potent feedback inhibition by glucosamine 6-phosphate . J Biol Chem 277 : 14764 – 14770 .

Brown ED Marquardt JL Lee JP Walsh CT Anderson KS ( 1994 ) Detection and characterization of a phospholactoyl-enzyme adduct in the reaction catalyzed by UDP- N -acetylglucosamine enolpyruvoyl transferase, MurZ . Biochemistry 33 : 10638 – 10645 .

Brown ED Vivas EI Walsh CT Kolter R ( 1995 ) MurA (MurZ), the enzyme that catalyzes the first committed step in peptidoglycan biosynthesis, is essential in Escherichia coli . J Bacteriol 177 : 4194 – 4197 .

Brown K Pompeo F Dixon S Mengin-Lecreulx D Cambillau C Bourne Y ( 1999 ) Crystal structure of the bifunctional N -acetylglucosamine 1-phosphate uridyltransferase from Escherichia coli : a paradigm for the related pyrophosphorylase superfamily . EMBO J 18 : 4096 – 4107 .

Bugg TDH Walsh CT ( 1992 ) Intracellular steps of bacterial cell wall peptidoglycan biosynthesis: enzymology, antibiotics, and antibiotic resistance . Nat Prod Rep 9 : 199 – 215 .

Bugg TDH Wright GD Dutka-Malen S Arthur M Courvalin P Walsh CT ( 1991 ) Molecular basis of vancomycin resistance in Enterococcus faecium BM4147: biosynthesis of depsipeptide peptidoglycan precursor by vancomycin resistance proteins VanH and VanA . Biochemistry 30 : 10408 – 10415 .

Burton E Gawande PV Yakandawala N LoVetri K Zhanel GG Romeo T Friesen AD Madhyastha S ( 2006 ) Antibiofilm activity of GlmU enzyme inhibitors against catheter-associated uropathogens . Antimicrob Agents Chemother 50 : 1835 – 1840 .

Byczynski B Mizyed S Berti PJ ( 2003 ) Nonenzymatic breakdown of the tetrahedral (α-carboxyketal phosphate) intermediates of MurA and AroA, two carboxyvinyl transferases. Protonation of different functional groups controls the rate and fate of breakdown . J Am Chem Soc 125 : 12541 – 12550 .

Candela T Fouet A ( 2006 ) Poly-gamma-glutamate in bacteria . Mol Microbiol 60 : 1091 – 1098 .

Chakravarty PK Greenlee WJ Parsons WH Patchett AA Combs P Roth A Busch RD Mellin TN ( 1989 ) (3-Amino-2-oxoalkyl)phosphonic acids and their analogues as novel inhibitors of d-alanine: d-alanine ligase . J Med Chem 32 : 1886 – 1890 .

Cheung KS Wasserman SA Dudek E Lerner SA Johnston M ( 1983 ) Chloralanyl and propargylglycyl dipeptides. Suicide substrate containing antibacterials . J Med Chem 26 : 1733 – 1741 .

Cheung KS Boisvert W Lerner SA Johnston M ( 1986 ) Chloroalanyl antibiotic peptides: antagonism of their antimicrobial effects by l-alanine and l-alanyl peptides in Gram-negative bacteria . J Med Chem 29 : 2060 – 2068 .

Chmara H Zähner H Borowski E ( 1984 ) Anticapsin, an active-site directed irreversible inhibitor of glucosamine 6-phosphate synthetase from Escherichia coli . J Antibiotics 37 : 1038 – 1043 .

Chmara H Andruszkiewicz R Borowski E ( 1985 ) Inactivation of glucosamine-6-phosphate synthetase from Salmonella typhimurium LT2 by fumaroyl diaminopropanoic acid derivatives, a novel group of glutamine analogs . Biochim Biophys Acta 870 : 357 – 366 .

Choi SY Esaki N Ashiuchi M Yoshimura T Soda K ( 1994 ) Bacterial glutamate racemase has high sequence similarity with myoglobins and forms an equimolar inactive complex with hemin . Proc Natl Acad Sci USA 91 : 10144 – 10147 .

Chopra I Storey C Falla JT Pearce JH ( 1998 ) Antibiotics, peptidoglycan synthesis and genomics: the chlamydial anomaly revisited . Microbiology 144 : 2673 – 2678 .

Comess KM Schurdak ME Voorbach MJ et al. ( 2006 ) An ultraefficient affinity-based high-throughout screening process: application to bacterial cell wall biosynthesis enzyme MurF . J Biomol Screen 11 : 743 – 754 .

Constantine KL Mueller L Goldfarb V Wittekind M Metzler WJ Yanchunas J Jr Robertson JG Malley MF Friedrichs MS Farmer BT II ( 1997 ) Characterization of NADP + binding to perdeuterated MurB: backbone atom NMR assignments and chemical-shift changes . J Mol Biol 267 : 1223 – 1246 .

Copié V Faraci SW Walsh CT Griffin RG ( 1988 ) Inhibition of alanine racemase by alanine phosphonate: detection of an imine linkage to pyridoxal 5′-phosphate in the enzyme-inhibitor complex by solid-state 15N nuclear magnetic resonance . Biochemistry 27 : 4966 – 4970 .

Cox RJ ( 1996 ) The DAP pathway of lysine as a target for antimicrobial agents . Nat Prod Rep 13 : 29 – 43 .

Cox RJ Sutherland A Vederas JC ( 2000 ) Bacterial diaminopimelate metabolism as a target for antibiotic design . Bioorg Med Chem 8 : 843 – 871 .

Dai HJ Parker CN Bao JJ ( 2002 ) Characterization and inhibition study of MurA enzyme by capillary electrophoresis . J Chromatogr B Analyt Technol Biomed Life Sci 766 : 123 – 132 .

Dai JB Liu Y Ray WJ Jr Konno M ( 1992 ) The crystal structure of muscle phosphoglucomutase refined at 2.7 angstrom resolution . J Biol Chem 267 : 6322 – 6337 .

Daniel RA Errington J ( 1993 ) DNA sequence of the murE - murD region of Bacillus subtilis 168 . J Gen Microbiol 139 : 361 – 370 .

Daub E Zawadzke LE Botstein D Walsh CT ( 1988 ) Isolation, cloning, and sequencing of the Salmonella typhimurium ddlA gene with purification and characterization of its product, d-alanine: d-alanine ligase (ADP forming) . Biochemistry 27 : 3701 – 3708 .

De Dios A Prieto L Martin JA et al. ( 2002 ) 4-Substituted d-glutamic acid analogues: the first potent inhibitors of glutamate racemase (MurI) enzyme with antibacterial activity . J Med Chem 45 : 4559 – 4570 .

De Smet KA Kempsell KE Gallagher A Duncan K Young DB ( 1999 ) Alteration of a single amino acid residue reverses fosfomycin resistance of recombinant MurA from Mycobacterium tuberculosis . Microbiology 145 : 3177 – 3184 .

Dementin S ( 2001 ) Etude du mécanisme réactionnel des Mur synthétases, enzymes impliquées dans la biosynthèse du peptidoglycane bactérien . Ph.D. Thesis, Université Paris-Sud, Orsay, France.

Dementin S Bouhss A Auger G Parquet C Mengin-Lecreulx D Dideberg O Van Heijenoort J Blanot D ( 2001 ) Evidence of a functional requirement for a carbamoylated lysine residue in MurD, MurE and MurF synthetases as established by chemical rescue experiments . Eur J Biochem 268 : 5800 – 5807 .

Deva T Baker EN Squire CJ Smith CA ( 2006 ) Structure of Escherichia coli UDP- N -acetylmuramoyl: l-alanine ligase (MurC) . Acta Cryst D62 : 1466 – 1474 .

DeVito JA Mills JA Liu VG et al. ( 2002 ) An array of target-specific screening strains for antibacterial discovery . Nat Biotechnol 20 : 478 – 483 .

Dhalla AM Yanchunas J Jr Ho HT Falk PJ Villafranca JJ Robertson JG ( 1995 ) Steady-state kinetic mechanism of Escherichia coli UDP- N -acetylenolpyruvylglucosamine reductase . Biochemistry 34 : 5390 – 5402 .

Dini C Drochon N Ferrari P Aszodi J ( 2000 ) Multi gram synthesis of UDP- N -acetylmuramic acid . Bioorg Med Chem Lett 10 : 143 – 145 .

Dodd D Reese JG Louer CR Ballard JD Spies MA Blanke SR ( 2007 ) Functional comparison of the two Bacillus anthracis glutamate racemases . J Bacteriol 189 : 5265 – 5275 .

Doublet P Van Heijenoort J Bohin J-P Mengin-Lecreulx D ( 1993 ) The murI gene of Escherichia coli is an essential gene that encodes a glutamate racemase activity . J Bacteriol 175 : 2970 – 2979 .

Doublet P Van Heijenoort J Mengin-Lecreulx D ( 1994 ) The glutamate racemase activity from Escherichia coli is regulated by peptidoglycan precursor UDP- N -acetylmuramoyl-l-alanine . Biochemistry 33 : 5285 – 5290 .

Doublet P Van Heijenoort J Mengin-Lecreulx D ( 1996 ) Regulation of the glutamate racemase of Escherichia coli investigated by site-directed mutagenesis . Microb Drug Resist 2 : 43 – 49 .

Du W Brown JR Sylvester DR Huang J Chalker AF So SY Holmes DJ Payne DJ Wallis NG ( 2000 ) Two active forms of UDP- N -acetylglucosamine enolpyruvyl transferase in Gram-positive bacteria . J Bacteriol 182 : 4146 – 4152 .

Duncan K Walsh CT ( 1988 ) ATP-dependent inactivation and slow binding inhibition of Salmonella typhimurium d-alanine: d-alanine ligase (ADP) by (aminoalkyl)phosphinate and aminophosphonate analogues of d-alanine . Biochemistry 27 : 3709 – 3714 .

Duncan K Faraci WS Matteson DS Walsh CT ( 1989 ) (1-Aminoethyl)boronic acid: a novel inhibitor for Bacillus stearothermophilus alanine racemase and Salmonella typhimuriumd-alanine: d-alanine ligase (ADP-forming) . Biochemistry 28 : 3541 – 3549 .

Duncan K Van Heijenoort J Walsh CT ( 1990 ) Purification and characterization of the d-alanyl-d-alanine-adding enzyme from Escherichia coli . Biochemistry 29 : 2379 – 2386 .

Ehmann DE Demeritt JE Hull KG Fisher SL ( 2004 ) Biochemical characterization of an inhibitor of Escherichia coli UDP- N -acetylmuramyl-l-alanine ligase . Biochim Biophys Acta 1698 : 167 – 174 .

Ellsworth BA Tom NJ Bartlett PA ( 1996 ) Synthesis and evaluation of inhibitors of bacterial d-alanine: d-alanine ligases . Chem Biol 3 : 37 – 44 .

El Zoeiby A Sanschagrin F Lamoureux J Darveau A Levesque RC ( 2000 ) Cloning, over-expression and purification of Pseudomonas aeruginosa murC encoding uridine diphosphate N -acetylmuramate: l-alanine ligase . FEMS Microbiol Lett 183 : 281 – 288 .

El Zoeiby A Sanschagrin F Havugimana PC Garnier A Levesque RC ( 2001 ) In vitro reconstruction of the biosynthetic pathway of peptidoglycan cytoplasmic precursor in Pseudomonas aeruginosa . FEMS Microbiol Lett 201 : 229 – 235 .

El Zoeiby A Sanschagrin F Levesque RC ( 2003a ) Structure and function of the Mur enzymes: development of novel inhibitors . Mol Microbiol 47 : 1 – 12 .

El Zoeiby A Sanschagrin F Darveau A Brisson J-R Levesque RC ( 2003b ) Identification of novel inhibitors of Pseudomonas aeruginosa MurC enzyme derived from phage-displayed peptide libraries . J Antimicrob Chemother 51 : 531 – 543 .

Emanuele JJ Jr Jin H Jacobson BL Chang CY Einspahr HM Villafranca JJ ( 1996 ) Kinetic and crystallographic studies of Escherichia coli UDP- N -acetylmuramate: l-alanine ligase . Prot Sci 5 : 2566 – 2574 .

Emanuele JJ Jr Jin H Yanchunas J Villafranca JJ ( 1997 ) Evaluation of the kinetic mechanism of Escherichia coli uridine diphosphate- N -acetylmuramate: l-alanine ligase . Biochemistry 36 : 7264 – 7271 .

Erion MD Walsh CT ( 1987 ) 1-Aminocyclopropanephosphonate: time-dependent inactivation of 1-aminocyclopropanecarboxylate deaminase and Bacillus stearothermophilus alanine racemase by slow dissociation behavior . Biochemistry 26 : 3417 – 3425 .

Esaki N Walsh CT ( 1986 ) Biosynthetic alanine racemase of Salmonella typhimurium : purification and characterization of the enzyme encoded by the alr gene . Biochemistry 25 : 3261 – 3267 .

Eschenburg S Schönbrunn E ( 2000 ) Comparative X-ray analysis of the un-liganded fosfomycin-target murA . Proteins 40 : 290 – 298 .

Eschenburg S Kabsch W Healy ML Schönbrunn E ( 2003 ) A new view of the mechanisms of UDP- N -acetylglucosamine enolpyruvyl transferase (MurA) and 5-enolpyruvylshikimate-3-phosphate synthase (AroA) derived from X-ray structures of their tetrahedral reaction intermediate states . J Biol Chem 278 : 49215 – 49222 .

Eschenburg S Priestman M Schönbrunn E ( 2005a ) Evidence that the fosfomycin target Cys 115 in UDP- N -acetylglucosamine enolpyruvyl transferase (MurA) is essential for product release . J Biol Chem 280 : 3757 – 3763 .

Eschenburg S Priestman MA Abdul-Latif FA Delachaume C Fassy FE ( 2005b ) A novel inhibitor that suspends the induced fit mechanism of UDP- N -acetylglucosamine enol pyruvyl transferase (MurA) . J Biol Chem 280 : 14070 – 14075 .

Eveland SS Pompliano DL Anderson MS ( 1997 ) Conditionally lethal Escherichia coli murein mutants contain point defects that map to regions conserved among murein and folyl poly-γ-glutamate ligases: identification of a ligase superfamily . Biochemistry 36 : 6223 – 6229 .

Falk PJ Ervin KM Volk KS Ho H-T ( 1996 ) Biochemical evidence for the formation of a covalent acyl-phosphate linkage between UDP- N -acetylmuramate and ATP in the Escherichia coli UDP- N -acetylmuramate: l-alanine ligase-catalyzed reaction . Biochemistry 35 : 1417 – 1422 .

Fan C Moews PC Walsh CT Knox JR ( 1994 ) Vancomycin resistance: structure of d-alanine: d-alanine ligase at 2.3 Å resolution . Science 266 : 439 – 443 .

Fan C Park IS Walsh CT Knox JR ( 1997 ) d-alanine: d-alanine ligase: phosphonate and phosphinate intermediates with wild type and the Y216F mutant . Biochemistry 36 : 2531 – 2538 .

Faraci WS Walsh CT ( 1989 ) Mechanism of inactivation of alanine racemase by β, β, β-trifluoroalanine . Biochemistry 28 : 431 – 437 .

Farmer BT II Constantine KL Goldfarb V Friedrichs MS Wittekind M Yanchunas J Jr Robertson JG Mueller L ( 1996 ) Localizing the NADP + binding site on the MurB enzyme by NMR . Nat Struct Biol 3 : 995 – 997 .

Fenn TD Stamper GF Morollo AA Ringe D ( 2003 ) A side reaction of alanine racemase: transamination of cycloserine . Biochemistry 42 : 5775 – 5783 .

Fenn TD Holyoak T Stamper GF Ringe D ( 2005 ) Effect of a Y265F mutant on the transamination-based cycloserine inactivation of alanine racemase . Biochemistry 44 : 5317 – 5327 .

Floquet N Mouilleron S Daher R Maigret B Badet B Badet-Denisot MA ( 2007a ) Ammonia channeling in bacterial glucosamine-6-phosphate synthase (Glms): molecular dynamics simulations and kinetic studies of protein mutants . FEBS Lett 581 : 2981 – 2987 .

Floquet N Richez C Durand P Maigret B Badet B Badet-Denisot MA ( 2007b ) Discovering new inhibitors of bacterial glucosamine-6P synthase (GlmS) by docking simulations . Bioorg Med Chem Lett 17 : 1966 – 1970 .

Flouret B Mengin-Lecreulx D Van Heijenoort J ( 1981 ) Reverse-phase high-pressure liquid chromatography of uridine diphosphate N -acetylmuramyl peptide precursors of bacterial cell wall peptidoglycan . Anal Biochem 114 : 59 – 63 .

Fotheringham IG Bledig SA Taylor PP ( 1998 ) Characterization of the genes encoding d-amino acid transaminase and glutamate racemase, two d-glutamate biosynthetic enzymes of Bacillus sphaericus ATCC 10208 . J Bacteriol 180 : 4319 – 4323 .

Francisco GD Li Z Albright JD et al. ( 2004 ) Phenyl thiazolyl urea and carbamate derivatives as new inhibitors of bacterial cell-wall biosynthesis . Bioorg Med Chem Lett 14 : 235 – 238 .

Gallo KA Knowles JR ( 1993 ) Purification, cloning, and cofactor independence of glutamate racemase from Lactobacillus . Biochemistry 32 : 3981 – 3990 .

Gallo KA Tanner ME Knowles JR ( 1993 ) Mechanism of the reaction catalyzed by glutamate racemase . Biochemistry 32 : 3991 – 3997 .

Galperin MY Koonin EV ( 1997 ) A diverse superfamily of enzymes with ATP-dependent carboxylate-amine/thiol ligase activity . Protein Sci 6 : 2639 – 2643 .

Gegnas LD Waddell ST Chabin RM Reddy S Wong KK ( 1998 ) Inhibitors of the bacterial cell wall biosynthesis enzyme MurD . Bioorg Med Chem Lett 8 : 1643 – 1648 .

Gehring AM Lees WJ Mindiola DJ Walsh CT Brown ED ( 1996 ) Acetyltransfer precedes uridylyltransfer in the formation of UDP- N -acetylglucosamine in separable active sites of the bifunctional GlmU protein of Escherichia coli . Biochemistry 35 : 579 – 585 .

Girardin SE Travassos LH Hervé M Blanot D Boneca IG Philpott DJ Sansonetti PJ Mengin-Lecreulx D ( 2003 ) Peptidoglycan molecular requirements allowing detection by Nod1 and Nod2 . J Biol Chem 278 : 41702 – 41708 .

Glanzmann P Gustafson J Komatsuzawa H Ohta K Berger-Bächi B ( 1999 ) glmM operon and methicillin-resistant glmM suppressor mutants in Staphylococcus aureus . Antimicrob Agents Chemother 43 : 240 – 245 .

Glavas S Tanner ME ( 1997 ) The inhibition of glutamate racemase by D- N -hydroxyglutamate . Bioorg Med Chem Lett 7 : 2265 – 2270 .

Glavas S Tanner ME ( 1999 ) Catalytic acid/base residues of glutamate racemase . Biochemistry 38 : 4106 – 4113 .

Glavas S Tanner ME ( 2001 ) Active site residues of glutamate racemase . Biochemistry 40 : 6199 – 6204 .

Gobec S Urleb U Auger G Blanot D ( 2001 ) Synthesis and biochemical evaluation of some novel N -acyl phosphono- and phosphinoalanine derivatives as potential inhibitors of the d-glutamic acid-adding enzyme . Pharmazie 56 : 295 – 297 .

Golinelli-Pimpaneau B Le Goffic F Badet B ( 1989 ) Glucosamine-6-phosphate synthase from Escherichia coli : mechanism of the reaction at the fructose 6-phosphate binding site . J Am Chem Soc 111 : 3029 – 3034 .

Gordon E Flouret B Chantalat L Van Heijenoort J Mengin-Lecreulx D Dideberg O ( 2001 ) Crystal structure of the UDP- N -acetylmuramoyl-l-alanyl-d-glutamate: meso-diaminopimelate ligase from Escherichia coli . J Biol Chem 276 : 10999 – 11006 .

Gu YG Florjancic AS Clark RF et al. ( 2004 ) Structure-activity relationships of novel potent MurF inhibitors . Bioorg Med Chem Lett 14 : 267 – 270 .

Gubler M Appoldt Y Keck W ( 1996 ) Overexpression, purification, and characterization of UDP- N -acetylmuramyl: l-alanine ligase from Escherichia coli . J Bacteriol 178 : 906 – 910 .

Hammes WP Neukam R Kandler O ( 1977 ) On the specificity of the uridine diphospho- N -acetylmuramyl-alanyl-d-glutamic acid: diamino acid ligase of Bifidobacterium globosum . Arch Microbiol 115 : 95 – 102 .

Healy VL Lessard IA Roper DI Knox JR Walsh CT ( 2000a ) Vancomycin resistance in enterococci: reprogramming of the d-Ala-d-Ala ligases in bacterial peptidoglycan biosynthesis . Chem Biol 7 : R109 – R119 .

Healy VL Mullins LS Li X Hall SE Raushel FM Walsh CT ( 2000b ) d-Ala-d-X ligases: evaluation of d-alanyl phosphate intermediate by MIX, PIX and rapid quench studies . Chem Biol 7 : 505 – 514 .

Hervé M Boniface A Gobec S Blanot D Mengin-Lecreulx D ( 2007 ) Biochemical characterization and physiological properties of Escherichia coli UDP- N -acetylmuramate: l-alanyl-γ-d-glutamyl-meso-diaminopimelate ligase (Mpl) . J Bacteriol 189 : 3987 – 3995 .

Hesse L Bostock J Dementin S Blanot D Mengin-Lecreulx D Chopra I ( 2003 ) Functional and biochemical analysis of Chlamydia trachomatis MurC, an enzyme displaying UDP- N -acetylmuramate: amino acid ligase activity . J Bacteriol 185 : 6507 – 6512 .

Heymann H Turdiu R Lee BK Barkulis SS ( 1968 ) A synthesis of uridinediphospho- N -acetylmuramic acid and its use as an acceptor of l-[ 14 C]alanine . Biochemistry 7 : 1393 – 1399 .

Hitchcock SA Eid CN Aikins JA Zia-Ebrahimi M Blaszczak LC ( 1998 ) The first total synthesis of bacterial cell wall precursor UDP- N -acetylmuramyl-pentapeptide (Park nucleotide) . J Am Chem Soc 120 : 1916 – 1917 .

Ho HT Falk PJ Ervin KM Krishnan BS Discotto LF Dougherty TJ Pucci MJ ( 1995 ) UDP- N -acetylmuramyl-l-alanine functions as an activator in the regulation of the Escherichia coli glutamate racemase activity . Biochemistry 34 : 2464 – 2470 .

Hofmann M Boles E Zimmermann FK ( 1994 ) Characterization of the essential yeast gene encoding N -acetylglucosamine-phosphate mutase . Eur J Biochem 221 : 741 – 747 .

Horton JR Bostock JM Chopra I Hesse L Phillips SE Adams DJ Johnson AP Fishwick CW ( 2003 ) Macrocyclic inhibitors of the bacterial cell wall biosynthesis enzyme MurD . Bioorg Med Chem Lett 13 : 1557 – 1560 .

Hove-Jensen B ( 1992 ) Identification of tms-26 as an allele of the gcaD gene, which encodes N -acetylglucosamine 1-phosphate uridyltransferase in Bacillus subtilis . J Bacteriol 174 : 6852 – 6856 .

Huber R Langworthy TA König H Thomm M Woese CR Sleytr UB Stetter KO ( 1986 ) Thermotoga maritima sp. nov. represents a new genus of unique extremely thermophylic eubacteria growing up to 90 °C . Arch Microbiol 144 : 324 – 333 .

Humljan J Kotnik M Boniface A Šolmajer T Urleb U Blanot D Gobec S ( 2006 ) A new approach towards peptidosulfonamides: synthesis of potential inhibitors of bacterial peptidoglycan biosynthesis enzymes MurD and MurE . Tetrahedron 62 : 10980 – 10988 .

Hutton CA Perugini MA Gerrard JA ( 2007 ) Inhibition of lysine biosynthesis: an evolving antibiotic strategy . Mol BioSyst 3 : 458 – 465 .

Huynh QK Gulve EA Dian T ( 2000 ) Purification and characterization of glutamine: fructose 6-phosphate amidotransferase from rat liver . Arch Biochem Biophys 379 : 307 – 313 .

Hwang KY Cho C-S Kim SS Sung H-C Yu YG Cho Y ( 1999 ) Structure and mechanism of glutamate racemase from Aquifex pyrophilus . Nat Struct Biol 6 : 422 – 426 .

Inagaki K Tanizawa K Badet B Walsh CT Tanaka H Soda K ( 1986 ) Thermostable alanine racemase from Bacillus stearothermophilus : molecular cloning of the gene, enzyme purification, and characterization . Biochemistry 25 : 3268 – 3274 .

Isupov MN Obmolova G Butterworth S Badet-Denisot MA Badet B Polikarpov I Littlechild JA Teplyakov A ( 1996 ) Substrate binding is required for assembly of the active conformation of the catalytic site in Ntn amidotransferases: evidence from the 1.8 Å crystal structure of the glutaminase domain of glucosamine 6-phosphate synthase . Structure 4 : 801 – 810 .

Jin H Emanuele JJ Jr Fairman R Robertson JG Hail ME Ho H-T Falk PJ Villafranca JJ ( 1996 ) Structural studies of Escherichia coli UDP- N -acetylmuramate: l-alanine ligase . Biochemistry 35 : 1423 – 1431 .

Jolly L Wu S Van Heijenoort J De Lencastre H Mengin-Lecreulx D Tomasz A ( 1997 ) The femR315 gene from Staphylococcus aureus , the interruption of which results in reduced methicillin resistance, encodes a phosphoglucosamine mutase . J Bacteriol 179 : 5321 – 5325 .

Jolly L Ferrari P Blanot D Van Heijenoort J Fassy F Mengin-Lecreulx D ( 1999 ) Reaction mechanism of phosphoglucosamine mutase from Escherichia coli . Eur J Biochem 262 : 202 – 210 .

Jolly L Pompeo F Van Heijenoort J Fassy F Mengin-Lecreulx D ( 2000 ) Autophosphorylation of phosphoglucosamine mutase from Escherichia coli . J Bacteriol 182 : 1280 – 1285 .

Katz AH Caufield CE ( 2003 ) Structure-based approaches to cell wall biosynthesis inhibitors . Curr Pharm Design 9 : 857 – 866 .

Kenig M Vandamme E Abraham EP ( 1976 ) The mode of action of bacilysin and anticapsin and biochemical properties of bacilysin-resistant mutants . J Gen Microbiol 94 : 46 – 54 .

Kim DH Lees WJ Walsh CT ( 1995 ) Stereochemical analysis of the tetrahedral adduct formed at the active site of UDP-GlcNAc enolpyruvyl transferase from the pseudosubstrates, ( E )- and ( Z )-fluorophosphoenolpyruvate, in D 2 O . J Am Chem Soc 117 : 6380 – 6381 .

Kim DH Lees WJ Kempsell KE Lane WS Duncan K Walsh CT ( 1996 ) Characterization of a Cys115 to Asp substitution in the Escherichia coli cell wall biosynthetic enzyme UDP-GlcNAc enolpyruvyl transferase (MurA) that confers resistance to inactivation by the antibiotic fosfomycin . Biochemistry 35 : 4923 – 4928 .

Kim K-H Bong Y-J Park JK Shin K-J Hwang KY Kim EE ( 2007 ) Structural basis for glutamate racamase inhibition . J Mol Biol 372 : 434 – 443 .

Kim MG Strych U Krause KL Benedik MJ Kohn H ( 2003a ) Evaluation of amino-substitued heterocyclic derivatives as alanine racemase inhibitors . Med Chem Res 12 : 130 – 138 .

Kim MG Strych U Krause K Benedik M Kohn H ( 2003b ) N (2)-substituted d,l-cycloserine derivatives: synthesis and evaluation as alanine racemase inhibitors . J Antibiot (Tokyo) 56 : 160 – 168 .

Kim MK Cho MK Song HE et al. ( 2007 ) Crystal structure of UDP- N -acetylenolpyruvylglucosamine reductase (MurB) from Thermus caldophilus . Proteins 66 : 751 – 754 .

Kim W-C Rhee H-I Park B-K Suk K-H Cha S-H ( 2000 ) Isolation of peptide ligands that inhibit glutamate racemase activity from a random phage display library . J Biomol Screen 5 : 435 – 440 .

Kimura K Tran LS Itoh Y ( 2004 ) Roles and regulation of the glutamate racemase isogenes, racE and yrpC , in Bacillus subtilis . Microbiology 150 : 2911 – 2920 .

Kobayashi K Ehrlich SD Albertini A et al. ( 2003 ) Essential Bacillus subtilis genes . Proc Natl Acad Sci USA 100 : 4678 – 4683 .

Kobayashi N Go N ( 1997 ) ATP binding proteins with different folds share a common ATP-binding structural motif . Nat Struct Biol 4 : 6 – 7 .

Kollonitsch J Barash L Kahan FM Kropp H ( 1973 ) New antibacterial agent via photofluorination of a bacterial cell wall constituent . Nature 243 : 346 – 347 .

Komatsuzawa H Fujiwara T Nishi H Yamada S Ohara M McCallum N Berger-Bächi B Sugai M ( 2004 ) The gate controlling cell wall synthesis in Staphylococcus aureus . Mol Microbiol 53 : 1221 – 1231 .

Kong D-X Zhu W-L Wu D-L Shen X Jiang H-L ( 2007 ) Comparison of 3D-QSAR methods using a novel class of MurF inhibitors . J Theor Comput Chem 6 : 63 – 80 .

Kostrewa D D'Arcy A Takacs B Kamber M ( 2001 ) Crystal structures of Streptococcus pneumoniae N -acetylglucosamine-1-phosphate uridyltransferase, GlmU, in apo form at 2.33 Å resolution and in complex with UDP- N -acetylglucosamine and Mg 2+ at 1.96 Å resolution . J Mol Biol 305 : 279 – 289 .

Kotnik M Oblak M Humljan J Gobec S Urleb U Šolmajer T ( 2004 ) Quantitative structure-activity relationships of Streptococcus pneumoniae MurD transition-state inhibitors . QSAR Comb Sci 23 : 399 – 405 .

Kotnik M Štefanič Anderluh P Preželj A ( 2007a ) Development of novel inhibitors targeting intracellular steps of peptidoglycan biosynthesis . Curr Pharm Des 13 : 2283 – 2309 .

Kotnik M Humljan J Contreras-Martel C et al. ( 2007b ) Structural and functional characterization of enantiomeric glutamic acid derivatives as potential transition state analogue inhibitors of MurD ligase . J Mol Biol 370 : 107 – 115 .

Kovač A Majce V Lenaršič R Bombek S Bostock JM Chopra I Polanc S Gobec S ( 2007 ) Diazenedicarboxamides as inhibitors of d-alanine-d-alanine ligase (Ddl) . Bioorg Med Chem Lett 17 : 2047 – 2054 .

Krekel F Samland AK Macheroux P Amrhein N Evans JN ( 2000 ) Determination of the p K a value of C115 in MurA (UDP- N -acetylglucosamine enolpyruvyltransferase) from Enterobacter cloacae . Biochemistry 39 : 12671 – 12677 .

Kurosu M Mahapatra S Narayanasamy P Crick DC ( 2007 ) Chemoenzymatic synthesis of Park's nucleotide: toward the development of high-throughput screening of MraY inhibitors . Tetrahedron Lett 48 : 799 – 803 .

Kutterer KM Davis JM Singh G Yang Y Hu W Severin A Rasmussen BA Krishnamurthy G Failli A Katz AH ( 2005 ) 4-Alkyl and 4,4′-dialkyl 1,2-bis(4-chlorophenyl)pyrazolidine-3,5-dione derivatives as new inhibitors of bacterial cell wall biosynthesis . Bioorg Med Chem Lett 15 : 2527 – 2531 .

Kuzin AP Sun T Jorczak-Baillass J Healy VL Walsh CT Knox JR ( 2000 ) Enzymes of vancomycin resistance: the structure of d-alanine-d-lactate ligase of naturally resistant Leuconostoc mesenteroides . Structure 8 : 463 – 470 .

Labaudiniere RF Xiang Y Jalluri RK Arvanites AC ( 2005 ) Antibiotic cycloalkyltetrahydroquinolone derivatives . US Patent, application number WO2005025556 A2.

Lacoste A-M Poulsen M Cassaigne A Neuzil E ( 1979 ) Inhibition of d-alanyl-d-alanine ligase in different bacterial species by amino phosphonic acids . Curr Microbiol 2 : 113 – 117 .

Lacoste A-M Chollet-Gravey A-M Vo Quang L Vo Quang Y Le Goffic F ( 1991 ) Time-dependent inhibition of Streptococcus faecalis d-alanine: d-alanine ligase by α-aminophosphonamidic acids . Eur J Med Chem 26 : 255 – 260 .

Lee JH Na Y Song HE et al. ( 2006a ) Crystal structure of the apo form of d-alanine: d-alanine ligase (Ddl) from Thermus caldophilus : a basis for the substrate-induced conformational changes . Proteins 64 : 1078 – 1082 .

Lee SG Hong SP Song JJ Kim SJ Kwak MS Sung MH ( 2006b ) Functional and structural characterization of thermostable d-amino acid aminotransferases from Geobacillus spp . Appl Environ Microbiol 72 : 1588 – 1594 .

Lees WJ Walsh CT ( 1995 ) Analysis of the enol ether transfer catalyzed by UDP-GlcNAc enolpyruvyl transferase using ( E )- and ( Z )-isomers of phosphoenolbutyrate: stereochemical, partitioning, and isotope effect studies . J Am Chem Soc 117 : 7329 – 7337 .

Lees WJ Benson TE Hogle JM Walsh CT ( 1996 ) ( E )-Enolbutyryl-UDP- N -acetylglucosamine as a mechanistic probe of UDP- N -acetylenolpyruvylglucosamine reductase (MurB) . Biochemistry 35 : 1342 – 1351 .

LeMagueres P Im H Ebalunode J Strych U Benedik MJ Briggs JM Kohn H Krause KL ( 2005 ) The 1.9 Å crystal structure of alanine racemase from Mycobacterium tuberculosis contains a conserved entryway into the active site . Biochemistry 44 : 1471 – 1481 .

Le Roux P Blanot D Mengin-Lecreulx D Van Heijenoort J ( 1991 ) Peptides containing 2-aminopimelic acid. Synthesis and study of in vitro effects on bacterial cells . Int J Peptide Protein Res 37 : 103 – 111 .

Le Roux P Auger G Van Heijenoort J Blanot D ( 1992 ) Synthesis of new peptide inhibitors of the meso -diaminopimelate-adding enzyme . Eur J Med Chem 27 : 899 – 907 .

Leung DK Andrews PR Craik DJ Iskander MN Winkler DA ( 1985 ) Design, synthesis and testing of transition state analogues of alanine racemase as antibacterials . Australian J Chem 38 : 297 – 206 .

Li Z Francisco GD Hu W et al. ( 2003 ) 2-Phenyl-5,6-dihydro-2 H -thieno[3,2- c ]pyrazol-3-ol derivatives as new inhibitors of bacterial cell wall biosynthesis . Bioorg Med Chem Lett 13 : 2591 – 2594 .

Liger D Blanot D Van Heijenoort J ( 1991 ) Effect of various alanine analogues on the l-alanine-adding enzyme from Escherichia coli . FEMS Microbiol Lett 80 : 111 – 116 .

Liger D Masson A Blanot D Van Heijenoort J Parquet C ( 1995 ) Over-production, purification and properties of the uridine-diphosphate- N -acetylmuramate: l-alanine ligase from Escherichia coli . Eur J Biochem 230 : 80 – 87 .

Liger D Masson A Blanot D Van Heijenoort J Parquet C ( 1996 ) Study of the overproduced uridine-diphosphate- N -acetylmuramate: l-alanine ligase from Escherichia coli . Microb Drug Resist 2 : 25 – 27 .

Liu H Sadamoto R Sears PS Wong C-H ( 2001 ) An efficient chemoenzymatic strategy for the synthesis of wild-type and vancomycin-resistant bacterial cell-wall precursors: UDP-N-acetylmuramyl-peptides . J Am Chem Soc 123 : 9916 – 9917 .

Liu S Chang JS Herberg JT Horng M-M Tomich PK Lin AH Marotti KR ( 2006 ) Allosteric inhibition of Staphylococcus aureus d-alanine: d-alanine ligase revealed by crystallographic studies . Proc Natl Acad Sci USA 103 : 15178 – 15183 .

Longenecker K Stamper GF Hajduk PJ et al. ( 2005 ) Structure of MurF from Streptococcus pneumoniae co-crystallized with a small molecule inhibitor exhibits interdomain closure . Protein Sci 14 : 3039 – 3047 .

Ludovice AM Wu SW De Lencastre H ( 1998 ) Molecular cloning and DNA sequencing of the Staphylococcus aureus UDP- N -acetylmuramyl tripeptide synthetase ( murE ) gene, essential for the optimal expression of methicillin resistance . Microb Drug Resist 4 : 85 – 90 .

Lundqvist T Fisher SL Kern G Folmer RH Xue Y Newton DT Keating TA Alm RA De Jonge BL ( 2007 ) Exploitation of structural and regulatory diversity in glutamate racemases . Nature 447 : 817 – 822 .

Mahapatra S Crick DC Brennan PJ ( 2000 ) Comparison of the UDP- N -acetylmuramate: l-alanine ligase enzymes from Mycobacterium tuberculosis and Mycobacterium leprae . J Bacteriol 182 : 6827 – 6830 .

Major DT Gao J ( 2006 ) A combined quantum mechanical and molecular mechanical study of the reaction mechanism and α-amino acidity in alanine racemase . J Am Chem Soc 128 : 16345 – 16357 .

Marmor S Petersen CP Reck F Yang W Gao N Fisher SL ( 2001 ) Biochemical characterization of a phosphinate inhibitor of Escherichia coli MurC . Biochemistry 40 : 12207 – 12214 .

Marquardt JL Siegele DA Kolter R Walsh CT ( 1992 ) Cloning and sequencing of Escherichia coli murZ and purification of its product, a UDP- N -acetylglucosamine enolpyruvyl transferase . J Bacteriol 174 : 5748 – 5752 .

Marquardt JL Brown ED Walsh CT Anderson KS ( 1993 ) Isolation and structural elucidation of a tetrahedral intermediate in the UDP- N -acetylglucosamine enolpyruvoyl transferase enzymatic pathway . J Am Chem Soc 115 : 10398 – 10399 .

Marquardt JL Brown ED Lane WS Haley TM Ichikawa Y Wong CH Walsh CT ( 1994 ) Kinetics, stoichiometry, and identification of the reactive thiolate in the inactivation of UDP-GlcNAc enolpyruvoyl transferase by the antibiotic fosfomycin . Biochemistry 33 : 10646 – 10651 .

Maruyama D Nishitani Y Nonaka T Kita A Fukami TA Mio T Yamada-Okabe H Yamada-Okabe T Miki K ( 2007 ) Crystal structure of uridine-diphospho- N -acetylglucosamine pyrophosphorylase from Candida albicans and catalytic reaction mechanism . J Biol Chem 282 : 17221 – 17230 .

May M Mehboob S Mulhearn DC Wang Z Yu H Thatcher GR Santarsiero BD Johnson ME Mesecar AD ( 2007 ) Structural and functional analysis of two glutamate racemase isozymes from Bacillus anthracis and implications for inhibitor design . J Mol Biol 371 : 1219 – 1237 .

McCoy AJ Maurelli AT ( 2005 ) Characterization of Chlamydia MurC-Ddl, a fusion protein exhibiting d-alanyl-d-alanine ligase activity involved in peptidoglycan synthesis and d-cycloserine sensitivity . Mol Microbiol 57 : 41 – 52 .

McCoy AJ Sandlin RC Maurelli AT ( 2003 ) In vitro and in vivo functional activity of Chlamydia MurA, a UDP- N -acetylglucosamine enolpyruvyl transferase involved in peptidoglycan synthesis and fosfomycin resistance . J Bacteriol 185 : 1218 – 1228 .

McDermott AE Creuzet F Griffin RG Zawadzke LE Ye Q-Z Walsh CT ( 1990 ) Rotational resonance determination of the structure of an enzyme-inhibitor complex: phosphorylation of an (aminoalkyl)phosphinate inhibitor of d-alanyl-d-alanine ligase by ATP . Biochemistry 29 : 5767 – 5775 .

Mengin-Lecreulx D Van Heijenoort J ( 1993 ) Identification of the glmU gene encoding N -acetylglucosamine-1-phosphate uridyltransferase in Escherichia coli . J Bacteriol 175 : 6150 – 6157 .

Mengin-Lecreulx D Van Heijenoort J ( 1994 ) Copurification of glucosamine-1-phosphate acetyltransferase and N -acetylglucosamine-1-phosphate uridyltransferase activities of Escherichia coli : characterization of the glmU gene product as a bifunctional enzyme catalyzing two subsequent steps in the pathway for UDP-N-acetylglucosamine synthesis . J Bacteriol 176 : 5788 – 5795 .

Mengin-Lecreulx D Van Heijenoort J ( 1996 ) Characterization of the essential gene glmM encoding phosphoglucosamine mutase in Escherichia coli . J Biol Chem 271 : 32 – 39 .

Mengin-Lecreulx D Flouret B Van Heijenoort J ( 1982 ) Cytoplasmic steps of peptidoglycan synthesis in Escherichia coli . J Bacteriol 151 : 1109 – 1117 .

Mengin-Lecreulx D Michaud C Richaud C Blanot D Van Heijenoort J ( 1988 ) Incorporation of ll-diaminopimelic acid into peptidoglycan of Escherichia coli mutants lacking diaminopimelate epimerase encoded by dapF . J Bacteriol 170 : 2031 – 2039 .

Mengin-Lecreulx D Parquet C Desviat LR Plá J Flouret B Ayala JA Van Heijenoort J ( 1989 ) Organization of the murE - murG region of Escherichia coli : identification of the murD gene encoding the d-glutamic-acid-adding enzyme . J Bacteriol 171 : 6126 – 6134 .

Mengin-Lecreulx D Blanot D Van Heijenoort J ( 1994 ) Replacement of diaminopimelic acid by cystathionine or lanthionine in the peptidoglycan of Escherichia coli . J Bacteriol 176 : 4321 – 4327 .

Mengin-Lecreulx D Van Heijenoort J Park JT ( 1996 ) Identification of the mpl gene encoding UDP- N -acetylmuramate: l-alanyl-γ-d-glutamyl-meso-diaminopimelate ligase in Escherichia coli and its role in recycling of cell wall peptidoglycan . J Bacteriol 178 : 5347 – 5352 .

Mengin-Lecreulx D Ayala J Bouhss A Van Heijenoort J Parquet C Hara H ( 1998 ) Contribution of the P mra promoter to expression of genes in the Escherichia coli mra cluster of cell envelope biosynthesis and cell division genes . J Bacteriol 180 : 4406 – 4412 .

Mengin-Lecreulx D Falla T Blanot D Van Heijenoort J Adams DJ Chopra I ( 1999 ) Expression of the Staphylococcus aureus UDP- N -acetylmuramoyl-l-alanyl-d-glutamate: l-lysine ligase in Escherichia coli and effects on peptidoglycan biosynthesis and cell growth . J Bacteriol 181 : 5909 – 5914 .

Michaud C Blanot D Flouret B Van Heijenoort J ( 1987 ) Partial purification and specificity studies of the d-glutamate-adding and d-alanyl-d-alanine-adding enzymes from Escherichia coli K12 . Eur J Biochem 166 : 631 – 637 .

Michaud C Mengin-Lecreulx D Van Heijenoort J Blanot D ( 1990 ) Over-production, purification and properties of the uridine-diphosphate- N -acetylmuramoyl-l-alanine-d-glutamate: meso-2,6-diaminopimelate ligase from Escherichia coli . Eur J Biochem 194 : 853 – 861 .

Milewski S Gabriel I Olchowy J ( 2006 ) Enzymes of UDP-GlcNAc biosynthesis in yeast . Yeast 23 : 1 – 14 .

Miller J Hammond SM Anderluzzi D Bugg TDH ( 1998 ) Aminoalkylphosphinate inhibitors of d-Ala-d-Ala adding enzyme . J Chem Soc, Perkin Trans 1 131 – 142 .

Mio T Yabe T Arisawa M Yamada-Okabe H ( 1998 ) The eukaryotic UDP- N -acetylglucosamine pyrophosphorylases: gene cloning, protein expression, and catalytic mechanism . J Biol Chem 273 : 14392 – 14397 .

Mio T Yamada-Okabe T Arisawa M Yamada-Okabe H ( 1999 ) Saccharomyces cerevisiae GNA1, an essential gene encoding a novel acetyltransferase involved in UDP- N -acetylglucosamine synthesis . J Biol Chem 274 : 424 – 429 .

Miyakawa T Matsuzawa H Matsuhashi M Sugino Y ( 1972 ) Cell wall peptidoglycan mutants of Escherichia coli K-12: existence of two clusters of genes, mra and mrb, for cell wall peptidoglycan biosynthesis . J Bacteriol 112 : 950 – 958 .

Mizyed S Oddone A Byczynski B Hughes DW Berti PJ ( 2005 ) UDP- N -acetylmuramic acid (UDP-MurNAc) is a potent inhibitor of MurA (enolpyruvyl-UDP-GlcNAc synthase) . Biochemistry 44 : 4011 – 4017 .

Möbitz H Bruice TC ( 2004 ) Multiple substrate binding states and chiral recognition in cofactor-independent glutamate racemase: a molecular dynamics study . Biochemistry 43 : 9685 – 9694 .

Moe ST Ala PJ Perola E ( 2003 ) Heterocyclic compounds and uses thereof as d -alanyl- d -alanine ligase inhibitors . US Patent, application number PCT/US02/20567, WO 03/001887 A2.

Mol CD Brooun A Dougan DR Hilgers MT Tari LW Wijnands RA Knuth MW McRee DE Swanson RW ( 2003 ) Crystal structures of active fully assembled substrate- and product-bound complexes of UDP- N -acetylmuramic acid: l-alanine ligase (MurC) from Haemophilus influenzae . J Bacteriol 185 : 4152 – 4162 .

Molina-López J Sanschagrin F Levesque RC ( 2006 ) A peptide inhibitor of MurA UDP- N -acetylglucosamine enolpyruvyl transferase: the first committed step in peptidoglycan biosynthesis . Peptides 27 : 3115 – 3121 .

Morollo AA Petsko GA Ringe D ( 1999 ) Structure of a Michaelis complex analogue: propionate binds in the substrate carboxylate site of alanine racemase . Biochemistry 38 : 3293 – 3301 .

Moulder JW ( 1993 ) Why is Chlamydia sensitive to penicillin in the absence of peptidoglycan? Infect Agents Dis 2 : 87 – 99 .

Mullins LS Zawadzke LE Walsh CT Raushel FM ( 1990 ) Kinetic evidence for the formation of d-alanyl phosphate in the mechanism of d-alanyl-d-alanine ligase . J Biol Chem 265 : 8993 – 8998 .

Mustata GI Briggs JM ( 2002 ) A structure-based design approach for the identification of novel inhibitors: application to an alanine racemase . J Comput Aided Mol Des 16 : 935 – 953 .

Nakajima N Tanizawa K Tanaka H Soda K ( 1986 ) Cloning and expression in Escherichia coli of the glutamate racemase gene from Pediococcus pentosaceus . Agric Biol Chem 50 : 2823 – 2830 .

Narayan RS VanNieuwenhze MS ( 2007 ) Synthesis of substrates and biochemical probes for study of the peptidoglycan biosynthetic pathway . Eur J Org Chem 1399 – 1414 .

Navia MA Ala PJ Griffith JP Ali JA Faerman CH Moe ST Magee AS Connelly PR Perola E ( 2003 ) Structure-based drug design methods for identifying d -Ala- d -Ala ligase inhibitors as antibacterial drugs . US Patent, application number PCT/US02/20465, WO 03/002063 A2.

Neuhaus FC ( 1962a ) The enzymatic synthesis of d-alanyl-dalanine. I. Purification and properties of d-alanyl-d-alanine synthetase . J Biol Chem 237 : 778 – 786 .

Neuhaus FC ( 1962b ) The enzymatic synthesis of d-alanyl-d-alanine. II. Kinetic studies on d-alanyl-d-alanine synthetase . J Biol Chem 237 : 3128 – 3135 .

Neuhaus FC Lynch JL ( 1964 ) The enzymatic synthesis of d-alanyl-d-alanine. III. On the inhibition of d-alanyl-d-alanine synthetase by the antibiotic d-cycloserine . Biochemistry 3 : 471 – 480 .

Neuhaus FC Struve WG ( 1965 ) Enzymatic synthesis of analogs of the cell-wall precursor. I. Kinetics and specificity of uridine diphospho- N -acetylmuramyl-l-alanyl-d-glutamyl-l-lysine :d-alanyl-d-alanine ligase (adenosine diphosphate) from Streptococcus faecalis R . Biochemistry 4 : 120 – 131 .

Neuhaus FC Hammes WP ( 1981 ) Inhibition of cell wall biosynthesis by analogues of alanine . Pharmac Ther 14 : 265 – 319 .

Nishida S Kurokawa K Matsuo M Sakamoto K Ueno K Kita K Sekimizu K ( 2006 ) Identification and characterization of amino acid residues essential for the active site of UDP- N -acetylenolpyruvylglucosamine reductase (MurB) from Staphylococcus aureus . J Biol Chem 281 : 1714 – 1724 .

Nishitani Y Maruyama D Nonaka T Kita A Fukami TA Mio T Yamada-Okabe H Yamada-Okabe T Miki K ( 2006 ) Crystal structures of N -acetylglucosamine-phosphate mutase, a member of the a-d-phosphohexomutase superfamily, and its substrate and product complexes . J Biol Chem 281 : 19740 – 19747 .

Nosal F Masson A Legrand R Blanot D Schoot B Van Heijenoort J Parquet C ( 1998 ) Site-directed mutagenesis and chemical modification of the two cysteine residues of the UDP- N -acetylmuramoyl: l-alanine ligase of Escherichia coli . FEBS Lett 426 : 309 – 313 .

Obmolova G Badet-Denisot MA Badet B Teplyakov A ( 1994 ) Crystallization and preliminary X-ray analysis of the two domains of glucosamine-6-phosphate synthase from Escherichia coli . J Mol Biol 242 : 703 – 705 .

Oikawa T Tauch A Schaffer S Fujioka T ( 2006 ) Expression of alr gene from Corynebacterium glutamicum ATCC 13032 in Escherichia coli and molecular characterization of the recombinant alanine racemase . J Biotechnol 125 : 503 – 512 .

Olsen LR Roderick SL ( 2001 ) Structure of the Escherichia coli GlmU pyrophosphorylase and acetyltransferase active sites . Biochemistry 40 : 1913 – 1921 .

Olsen LR Vetting MW Roderick SL ( 2007 ) Structure of the E. coli bifunctional GlmU acetyltransferase active site with substrates and products . Protein Sci 16 : 1230 – 1235 .

Paradis-Bleau C Beaumont M Boudreault L Lloyd A Sanschagrin F Bugg TD Levesque RC ( 2006 ) Selection of peptide inhibitors against the Pseudomonas aeruginosa MurD cell wall enzyme . Peptides 27 : 1693 – 1700 .

Park JT ( 1952 ) Uridine 5′-pyrophosphate derivatives. I. Isolation from Staphylococcus aureus . J Biol Chem 194 : 877 – 884 .

Parsons WH Patchett AA Bull HG et al. ( 1988 ) Phosphinic acid inhibitors of d-alanyl-d-alanine ligase . J Med Chem 31 : 1772 – 1778 .

Patchett AA Taub D Weissberger B Valiant ME Gadebusch H Thornberry NA Bull HG ( 1988 ) Antibacterial activities of fluorovinyl- and chlorovinylglycine and several derived dipeptides . Antimicrob Agents Chemother 32 : 319 – 323 .

Patte J-C ( 1983 ) Diaminopimelate and lysine . Amino Acids: Biosynthesis and Genetic Regulation ( Herrmann KL & Somerville RL , eds), pp. 213 – 228 . Addison-Wesley Publishing Co. , Reading, MA.

Peisach D Chipman DM Van Ophem PW Manning JM Ringe D ( 1998 ) Crystallographic study of steps along the reaction pathway of d-amino acid aminotransferase . Biochemistry 37 : 4958 – 4967 .

Peneff C Mengin-Lecreulx D Bourne Y ( 2001a ) The crystal structures of Apo and complexed Saccharomyces cerevisiae GNA1 shed light on the catalytic mechanism of an amino-sugar N -acetyltransferase . J Biol Chem 276 : 16328 – 16334 .

Peneff C Ferrari P Charrier V Taburet Y Monnier C Zamboni V Winter J Harnois M Fassy F Bourne Y ( 2001b ) Crystal structures of two human pyrophosphorylase isoforms in complexes with UDPGlc(Gal)NAc: role of the alternatively spliced insert in the enzyme oligomeric assembly and active site architecture . EMBO J 20 : 6191 – 6202 .

Perdih A Kotnik M Hodošček M Šolmajer T ( 2007 ) Targeted molecular dynamics simulation studies of binding and conformational changes in E. coli MurD . Proteins 68 : 243 – 254 .

Pompeo F Van Heijenoort J Mengin-Lecreulx D ( 1998 ) Probing the role of cysteine residues in glucosamine-1-phosphate acetyltransferase activity of the bifunctional GlmU protein from Escherichia coli : site-directed mutagenesis and characterization of the mutant enzymes . J Bacteriol 180 : 4799 – 4803 .

Pompeo F Bourne Y Van Heijenoort J Fassy F Mengin-Lecreulx D ( 2001 ) Dissection of the bifunctional Escherichia coli N -acetylglucosamine-1-phosphate uridyltransferase enzyme into autonomously functional domains and evidence that trimerization is absolutely required for glucosamine-1-phosphate acetyltransferase activity and cell growth . J Biol Chem 276 : 3833 – 3839 .

Pratviel-Sosa F Mengin-Lecreulx D Van Heijenoort J ( 1991 ) Over-production, purification and properties of the uridine diphosphate N -acetylmuramoyl-l-alanine: d-glutamate ligase from Escherichia coli . Eur J Biochem 202 : 1169 – 1176 .

Pratviel-Sosa F Acher F Trigalo F Blanot D Azerad R Van Heijenoort J ( 1994 ) Effect of various analogues of d-glutamic acid on the d-glutamate-adding enzyme from Escherichia coli . FEMS Microbiol Lett 115 : 223 – 228 .

Pucci MJ Thanassi JA Ho H-T Falk PJ Dougherty TJ ( 1995 ) Staphylococcus haemolyticus contains two d-glutamic acid biosynthetic activities, a glutamate racemase and a d-amino acid transaminase . J Bacteriol 177 : 336 – 342 .

Pucci MJ Thanassi JA Discotto LF Kessler RE Dougherty TJ ( 1997 ) Identification and characterization of cell wall-cell division gene clusters in pathogenic Gram-positive cocci . J Bacteriol 179 : 5632 – 5635 .

Puig E Garcia-Viloca M González-Lafont A López I Daura X Lluch JM ( 2005 ) A molecular dynamics simulation of the binding modes of d-glutamate and d-glutamine to glutamate racemase . J Chem Theory Comput 1 : 737 – 749 .

Puig E Garcia-Viloca M González-Lafont A Lluch JM ( 2006 ) On the ionization state of the substrate in the active site of glutamate racemase. A QM/MM study about the importance of being zwitterionic . J Phys Chem A 110 : 717 – 725 .

Raczynska J Olchowy J Konariev PV Svergun DI Milewski S Rypniewski W ( 2007 ) The crystal and solution studies of glucosamine-6-phosphate synthase from Candida albicans . J Mol Biol 372 : 672 – 688 .

Raetz CR Roderick SL ( 1995 ) A left-handed parallel beta helix in the structure of UDP- N -acetylglucosamine acyltransferase . Science 270 : 997 – 1000 .

Ramilo C Appleyard RJ Wanke C Krekel F Amrhein N Evans JN ( 1994 ) Detection of the covalent intermediate of UDP- N -acetylglucosamine enolpyruvyl transferase by solution-state and time-resolved solid-state NMR spectroscopy . Biochemistry 33 : 15071 – 15079 .

Ramos-Aires J Plésiat P Kocjancic-Curty L Köhler T ( 2004 ) Selection of an antibiotic-hypersusceptible mutant of Pseudomonas aeruginosa : identification of the GlmR transcriptional regulator . Antimicrob Agents Chemother 48 : 843 – 851 .

Raymond JB Price NP Pavelka MS Jr ( 2003 ) A method for the enzymatic synthesis and HPLC purification of the peptidoglycan precursor UDP- N -acetylmuramic acid . FEMS Microbiol Lett 229 : 83 – 89 .

Reck F Marmor S Fisher S Wuonola MA ( 2001 ) Inhibitors of the bacterial cell wall biosynthesis enzyme MurC . Bioorg Med Chem Lett 11 : 1451 – 1454 .

Reddy S Waddell ST Kuo DW Wong KK Pompliano DL ( 1999 ) Preparative enzymatic synthesis and characterization of the cytoplasmic intermediates of murein biosynthesis . J Am Chem Soc 121 : 1175 – 1178 .

Richaud C Mengin-Lecreulx D Pochet S Johnson EJ Cohen GN Marlière P ( 1993 ) Directed evolution of biosynthetic pathways: recruitment of cysteine thioethers for constructing the cell wall of Escherichia coli . J Biol Chem 268 : 26827 – 26835 .

Rios A Amyes TL Richard JP ( 2000 ) Formation and stability of organic zwitterions in aqueous solution: enolates of the amino acid glycine and its derivatives . J Am Chem Soc 122 : 9373 – 9385 .

Rogers HJ Perkins HR Ward JB ( 1980 ) Microbial Cell Walls and Membranes . Chapman and Hall , London, UK.

Roper DI Huyton T Vagin A Dodson G ( 2000 ) The molecular basis of vancomycin resistance in clinically relevant Enterococci : crystal structure of d-alanyl-d-lactate ligase (VanA) . Proc Natl Acad Sci USA 97 : 8921 – 8925 .

Ruzheinikov SN Taal MA Sedelnikova SE Baker PJ Rice DW ( 2005 ) Substrate-induced conformational changes in Bacillus subtilis glutamate racemase and their implications for drug discovery . Structure 13 : 1707 – 1713 .

Saito M Nishimura K Hasegawa Y Shinohara T Wakabayashi S Kurihara T Ishizuka M Nagata Y ( 2007 ) Alanine racemase from Helicobacter pylori NCTC 11637: purification, characterization and gene cloning . Life Sci 80 : 788 – 794 .

Samland AK Amrhein N Macheroux P ( 1999 ) Lysine 22 in UDP- N -acetylglucosamine enol pyruvyl transferase from Enterobacter cloacae is crucial for enzymatic activity and the formation of covalent adducts with the substrate phospho enol pyruvate and the antibiotic fosfomycin . Biochemistry 38 : 13162 – 13169 .

Samland AK Etezady-Esfarjani T Amrhein N Macheroux P ( 2001 ) Asparagine 23 and aspartate 305 are essential residues in the active site of UDP- N -acetylglucosamine enol pyruvyl transferase from Enterobacter cloacae . Biochemistry 40 : 1550 – 1559 .

Sarver DW Rogers JM Epps DM ( 2002 ) Determination of ligand-MurB interactions by isothermal denaturation: application as a secondary assay to complement high throughput screening . J Biomol Screen 7 : 21 – 28 .

Sato M Kirimura K Kino K ( 2005 ) d-Amino acid dipeptide production utilizing d-alanine-d-alanine ligases with novel substrate specificity . J Biosci Bioeng 99 : 623 – 628 .

Sato M Kirimura K Kino K ( 2006 ) Substrate specificity of thermostable d-alanine-d-alanine ligase from Thermotoga maritima ATCC 43589 . Biosci Biotechnol Biochem 70 : 2790 – 2792 .

Sauvage E Charlier P Terrak M Ayala JA ( 2008 ) The structure of Penicillin-Binding Proteins and their role in peptidoglycan biosynthesis . FEMS Microbiol Rev , doi: DOI: 10.1111/j.1574-6976.2008.00105.x

Sawada S Tanaka Y Hayashi S Ryu M Hasegawa T Yamamoto Y Esaki N Soda K Takahashi S ( 1994 ) Kinetics of thermostable alanine racemase of Bacillus stearothermophilus . Biosci Biotechnol Biochem 58 : 807 – 811 .

Schleifer KH Kandler O ( 1972 ) Peptidoglycan types of bacterial cell walls and their taxonomic implications . Bacteriol Rev 36 : 407 – 477 .

Schönbrunn E Sack S Eschenburg S Perrakis A Krekel F Amrhein N Mandelkow E ( 1996 ) Crystal structure of UDP- N -acetylglucosamine enolpyruvyltransferase, the target of the antibiotic fosfomycin . Structure 4 : 1065 – 1075 .

Schönbrunn E Svergun DI Amrhein N Koch MH ( 1998 ) Studies on the conformational changes in the bacterial cell wall biosynthetic enzyme UDP- N -acetylglucosamine enol pyruvyltransferase (MurA) . Eur J Biochem 253 : 406 – 412 .

Schönbrunn E Eschenburg S Luger K Kabsch W Amrhein N ( 2000a ) Structural basis for the interaction of the fluorescence probe 8-anilino-1-naphthalene sulfonate (ANS) with the antibiotic target MurA . Proc Natl Acad Sci USA 97 : 6345 – 6349 .

Schönbrunn E Eschenburg S Krekel F Luger K Amrhein N ( 2000b ) Role of the loop containing residue 115 in the induced-fit mechanism of the bacterial cell wall biosynthetic enzyme MurA . Biochemistry 39 : 2164 – 2173 .

Schouten JA Bagga S Lloyd AJ De Pascale G Dowson CG Roper DI Bugg TD ( 2006 ) Fluorescent reagents for in vitro studies of lipid-linked steps of bacterial peptidoglycan biosynthesis: derivatives of UDPMurNAc-pentapeptide containing d-cysteine at position 4 or 5 . Mol Biosyst 2 : 484 – 491 .

Seebeck FP Hilvert D ( 2003 ) Conversion of a PLP-dependent racemase into an aldolase by a single active site mutation . J Am Chem Soc 125 : 10158 – 10159 .

Sengupta S Shah M Nagaraja V ( 2006 ) Glutamate racemase from Mycobacterium tuberculosis inhibits DNA gyrase by affecting its DNA-binding . Nucleic Acids Res 34 : 5567 – 5576 .

Seow TK Inagaki K Tamura T Soda K Tanaka H ( 1998 ) Alanine racemase from an acidophile, Acidiphilium organovorum : purification and characterisation . Biosci Biotechnol Biochem 62 : 242 – 247 .

Seow TK Inagaki K Nakamura T Maeda R Tamura T Tanaka H ( 2000 ) Purification and some characteristics of a monomeric alanine racemase from an extreme thermophile, Thermus thermophilus . J Biosci Bioeng 90 : 344 – 346 .

Shatalin KY Neyfakh AA ( 2005 ) Efficient gene inactivation in Bacillus anthracis . FEMS Microbiol Lett 245 : 315 – 319 .

Shaw JP Petsko GA Ringe D ( 1997 ) Determination of the structure of alanine racemase from Bacillus stearothermophilus at 1.9-Å resolution . Biochemistry 36 : 1329 – 1342 .

Sheng Y Sun X Shen Y Bognar AL Baker EN Smith CA ( 2000 ) Structural and functional similarities in the ADP-forming amide bond ligase superfamily: implications for a substrate-induced conformational change in folylpolyglutamate synthetase . J Mol Biol 302 : 427 – 440 .

Sim MM Ng SB Buss AD Crasta SC Goh KL Lee SK ( 2002 ) Benzylidene rhodanines as novel inhibitors of UDP- N -acetylmuramate/l-alanine ligase . Bioorg Med Chem Lett 12 : 697 – 699 .

Skarzynski T Mistry A Wonacott A Hutchinson SE Kelly VA Duncan K ( 1996 ) Structure of UDP- N -acetylglucosamine enolpyruvyl transferase, an enzyme essential for the synthesis of bacterial peptidoglycan, complexed with substrate UDP- N -acetylglucosamine and the drug fosfomycin . Structure 4 : 1465 – 1474 .

Skarzynski T Kim DH Lees WJ Walsh CT Duncan K ( 1998 ) Stereochemical course of enzymatic enolpyruvyl transfer and catalytic conformation of the active site revealed by the crystal structure of the fluorinated analogue of the reaction tetrahedral intermediate bound to the active site of the C115A mutant of MurA . Biochemistry 37 : 2572 – 2577 .

Smith CA ( 2006 ) Structure, function and dynamics in the mur family of bacterial cell wall ligases . J Mol Biol 362 : 640 – 655 .

Snyder NJ Tebbe MJ Victor F et al. ( 1999 ) Synthesis and evaluation of Streptococcus pneumoniae MurD transition state inhibitors . Abstracts of the 39th Interscience Conference on Antimicrobial Agents and Chemotherapy , September 26–29, San Francisco, USA, p. 330.

Spraggon G Schwarzenbacher R Kreusch A et al. ( 2004 ) Crystal structure of an Udp-N-acetylmuramate-alanine ligase MurC (TM0231) from Thermotoga maritima at 2.3 Å resolution . Proteins 55 : 1078 – 1081 .

Stamper GF Morollo AA Ringe D ( 1998 ) Reaction of alanine racemase with 1-aminoethylphosphonic acid forms a stable external aldimine . Biochemistry 37 : 10438 – 10445 .

Stamper GF Longenecker K Fry EH et al. ( 2006 ) Structure-based optimization of MurF inhibitors . Chem Biol Drug Res 67 : 58 – 65 .

Štrancar K Blanot D Gobec S ( 2006 ) Design, synthesis and structure-activity relationships of new phosphinate inhibitors of MurD . Bioorg Med Chem Lett 16 : 343 – 348 .

Štrancar K Boniface A Blanot D Gobec S ( 2007 ) Phosphinate inhibitors of UDP- N -acetylmuramoyl-l-alanyl-d-glutamate: l-lysine ligase (MurE) . Arch Pharm Chem Life Sci 340 : 127 – 134 .

Strominger JL Ito E Threnn RH ( 1960 ) Competitive inhibition of enzymatic reactions by oxamycin . J Am Chem Soc 82 : 998 – 999 .

Strych U Benedik MJ ( 2002 ) Mutant analysis shows that alanine racemases from Pseudomonas aeruginosa and Escherichia coli are dimeric . J Bacteriol 184 : 4321 – 4325 .

Strych U Huang HC Krause KL Benedik MJ ( 2000 ) Characterization of the alanine racemases from Pseudomonas aeruginosa PAO1 . Curr Microbiol 41 : 290 – 294 .

Strych U Penland RL Jimenez M Krause KL Benedik MJ ( 2001 ) Characterization of the alanine racemases from two mycobacteria . FEMS Microbiol Lett 196 : 93 – 98 .

Sugio S Petsko GA Manning JM Soda K Ringe D ( 1995 ) Crystal structure of a d-amino acid aminotransferase: how the protein controls stereoselectivity . Biochemistry 34 : 9661 – 9669 .

Sugio S Kashima A Kishimoto K Peisach D Petsko GA Ringe D Yoshimura T Esaki N ( 1998 ) Crystal structures of L201A mutant of d-amino acid aminotransferase at 2.0 Å resolution: implication of the structural role of Leu201 in transamination . Protein Eng 11 : 613 – 619 .

Sulzenbacher G Gal L Peneff C Fassy F Bourne Y ( 2001 ) Crystal structure of Streptococcus pneumoniae N -acetylglucosamine-1-phosphate uridyltransferase bound to acetyl-coenzyme A reveals a novel active site architecture . J Biol Chem 276 : 11844 – 11851 .

Sun S Toney MD ( 1999 ) Evidence for a two-base mechanism involving tyrosine-265 from arginine-219 mutants of alanine racemase . Biochemistry 38 : 4058 – 4065 .

Sun X Cross JA Bognar AL Baker EN Smith CA ( 2001 ) Folate-binding triggers the activation of folylpolyglutamate synthetase . J Mol Biol 310 : 1067 – 1078 .

Szumilo T Zeng Y Pastuszak I Drake R Szumilo H Elbein AD ( 1996 ) Purification to homogeneity and properties of UDP-GlcNAc (GalNAc) pyrophosphorylase . J Biol Chem 271 : 13147 – 13154 .

Tanizawa K Masu Y Asano S Tanaka H Soda K ( 1989 ) Thermostable d-amino acid aminotransferase from a thermophilic Bacillus species. Purification, characterization, and active site sequence determination . J Biol Chem 264 : 2445 – 2449 .

Tanner ME ( 2002 ) Understanding nature's strategies for enzyme-catalyzed racemization and epimerization . Acc Chem Res 35 : 237 – 246 .

Tanner ME Miao S ( 1994 ) The synthesis and stability of aziridino-glutamate, an irreversible inhibitor of glutamate racemase . Tetrahedron Lett 35 : 4073 – 4076 .

Tanner ME Vaganay S Van Heijenoort J Blanot D ( 1996 ) Phosphinate inhibitors of the d-glutamic acid-adding enzyme of peptidoglycan biosynthesis . J Org Chem 61 : 1756 – 1760 .

Tavares IM Jolly L Pompeo F Leitão JH Fialho AM Sá-Correia I Mengin-Lecreulx D ( 2000 ) Identification of the Pseudomonas aeruginosa glmM gene, encoding phosphoglucosamine mutase . J Bacteriol 182 : 4453 – 4457 .

Tayeh MA Dotson GD Clemens JC Woodard RW ( 1995 ) Overproduction and one-step purification of Escherichia coli UDP- N -acetylglucosamine enolpyruvyl reductase . Protein Expr Purif 6 : 757 – 762 .

Taylor PP Fortheringham IG ( 1997 ) Nucleotide sequence of the Bacillus licheniformis ATTC 10716 dat gene and comparison of the predicted amino acid sequence with those of other bacterial species . Biochim Biophys Acta 1350 : 38 – 40 .

Teplyakov A Obmolova G Badet-Denisot MA Badet B Polikarpov I ( 1998 ) Involvement of the C terminus in intramolecular nitrogen channeling in glucosamine 6-phosphate synthase: evidence from a 1.6 Å crystal structure of the isomerase domain . Structure 6 : 1047 – 1055 .

Teplyakov A Obmolova G Badet B Badet-Denisot MA ( 2001 ) Channeling of ammonia in glucosamine-6-phosphate synthase . J Mol Biol 313 : 1093 – 1102 .

Thompson RJ Bouwer HG Portnoy DA Frankel FR ( 1998 ) Pathogenicity and immunogenicity of a Listeria monocytogenes strain that requires d-alanine for growth . Infect Immun 66 : 3552 – 3561 .

Thornberry NA Bull HG Taub D Greenlee WJ Patchett AA Cordes EH ( 1987 ) 3-Halovinylglycines. Efficient irreversible inhibitors of E. coli alanine racemase . J Am Chem Soc 109 : 7543 – 7544 .

Thornberry NA Bull HG Taub D Wilson KE Gimenez-Gallego G Rosegay A Soderman DD Patchett AA ( 1991 ) Mechanism-based inactivation of alanine racemase by 3-halovinylglycines . J Biol Chem 266 : 21657 – 21665 .

Traxinger RR Marshall S ( 1991 ) Coordinated regulation of glutamine: fructose-6-phosphate amidotransferase activity by insulin, glucose, and glutamine. Role of hexosamine biosynthesis in enzyme regulation . J Biol Chem 266 : 10148 – 10154 .

Ueda T Feng F Sadamoto R Niikura K Monde K Nishimura S-I ( 2004 ) Synthesis of 4-fluorinated UDP-MurNAc pentapeptide as an inhibitor of bacterial growth . Org Lett 6 : 1753 – 1756 .

Vaara M ( 1992 ) Eight bacterial proteins, including UDP- N -acetylglucosamine acyltransferase (LpxA) and three other transferases of Escherichia coli , consist of a six-residue periodicity theme . FEMS Microbiol Lett 76 : 249 – 254 .

Vaganay S Tanner ME Van Heijenoort J Blanot D ( 1996 ) Study of the reaction mechanism of the d-glutamic acid-adding enzyme from Escherichia coli . Microb Drug Resist 2 : 51 – 54 .

Van Assche I Soroka M Haemers A Hooper M Blanot D Van Heijenoort J ( 1991 ) Synthesis and antiobacterial evaluation of phosphonic acid analogues of diaminopimelic acid . Eur J Med Chem 26 : 505 – 515 .

Van Heijenoort J ( 2001 ) Recent advances in the formation of the bacterial peptidoglycan monomer subunit . Nat Prod Rep 18 : 503 – 519 .

Van Ophem PW Peisach D Erickson SD Soda K Ringe D Manning JM ( 1999 ) Effects of the E177K mutation in d-amino acid transaminase. Studies on an essential coenzyme anchoring group that contributes to stereochemical fidelity . Biochemistry 38 : 1323 – 1331 .

Victor F Tebbe MJ Birch GM Smith MC Letourneau DL Wu CE ( 1999 ) d -Glutamic acid analogs as Streptococcus pneumoniae UDP- N -acetyl muramoyl-alanine: d -glutamate ligase (MurD) inhibitors. Abstracts of the 39th Interscience Conference on Antimicrobial Agents and Chemotherapy , September 26–29, San Francisco, USA, p. 330.

Volkman BF Zhang Q Debabov DV Rivera E Kresheck GC Neuhaus FC ( 2001 ) Biosynthesis of d-alanyl-lipoteichoic acid: the tertiary structure of apo-d-alanyl carrier protein . Biochemistry 40 : 7964 – 7972 .

Vollmer W Blanot D De Pedro MA ( 2008 ) Peptidoglycan structure and architecture . FEMS Microbiol Rev , doi: DOI: 10.1111/j.1574-6976.2007.00094.x

Vo-Quang Y Carniato D Vo-Quang L Lacoste AM Neuzil E Le Goffic F ( 1986a ) (β-Chloro-α-aminoethyl)phosphonic acids as inhibitors of alanine racemase and d-alanine :d-alanine ligase . J Med Chem 29 : 148 – 151 .

Vo-Quang Y Carniato D Vo-Quang L Lacoste AM Neuzil E Le Goffic F ( 1986b ) (1-Amino-2-propenyl) phosphonic acid, an inhibitor of alanine racemase and d-alanine: d-alanine ligase . J Med Chem 29 : 579 – 581 .

Walker B Brown MF Lynas JF Martin SL McDowell A Badet B Hill AJ ( 2000 ) Inhibition of Escherichia coli glucosamine synthetase by novel electrophilic analogues of glutamine – comparison with 6-diazo-5-oxo-norleucine . Bioorg Med Chem Lett 10 : 2795 – 2798 .

Walsh CT ( 1989 ) Enzymes of the d-alanine branch of bacterial cell wall peptidoglycan assembly . J Biol Chem 264 : 2393 – 2396 .

Walsh CT Benson TE Kim DH Lees WJ ( 1996 ) The versatility of phosphoenolpyruvate and its vinyl ether products in biosynthesis . Chem Biol 3 : 83 – 91 .

Walsh AW Falk PJ Thanassi J Discotto L Pucci MJ Ho H-T ( 1999 ) Comparison of the d-glutamate-adding enzymes from selected Gram-positive and Gram-negative bacteria . J Bacteriol 181 : 5395 – 5401 .

Wang E Walsh C ( 1978 ) Suicide substrates for the alanine racemase of Escherichia coli B . Biochemistry 17 : 1313 – 1321 .

Wang EA Walsh C ( 1981 ) Characteristics of β,β-difluoroalanine and β,β,β-trifluoroalanine as suicide substrates of Escherichia coli B alanine racemase . Biochemistry 20 : 7539 – 7546 .

Wanke C Amrhein N ( 1993 ) Evidence that the reaction of the UDP- N -acetylglucosamine 1-carboxyvinyltransferase proceeds through the O -phosphothioketal of pyruvic acid bound to Cys115 of the enzyme . Eur J Biochem 218 : 861 – 870 .

Wanke C Falchetto R Amrhein N ( 1992 ) The UDP- N -acetylglucosamine 1-carboxyvinyl-transferase of Enterobacter cloacae . Molecular cloning, sequencing of the gene and overexpression of the enzyme . FEBS Lett 301 : 271 – 276 .

Ward JB ( 1984 ) Biosynthesis of peptidoglycan: points of attack of wall inhibitors . Pharmac Ther 25 : 327 – 369 .

Wasserman SA Daub E Grisafi P Botstein D Walsh CT ( 1984 ) Catabolic alanine racemase from Salmonella typhimurium : DNA sequence, enzyme purification, and characterization . Biochemistry 23 : 5182 – 5187 .

Watanabe A Yoshimura T Mikami B Esaki N ( 1999a ) Tyrosine 265 of alanine racemase serves as a base abstracting α-hydrogen from l-alanine: the counterpart residue to lysine 39 specific to d-alanine . J Biochem (Tokyo) 126 : 781 – 786 .

Watanabe A Kurokawa Y Yoshimura T Kurihara T Soda K Esaki N ( 1999b ) Role of lysine 39 of alanine racemase from Bacillus stearothermophilus that binds pyridoxal 5′-phosphate. Chemical rescue studies of Lys 39 →Ala mutant . J Biol Chem 274 : 4189 – 4194 .

Watanabe A Yoshimura T Mikami B Hayashi H Kagamiyama H Esaki N ( 2002 ) Reaction mechanism of alanine racemase from Bacillus stearothermophilus : X-ray crystallographic studies of the enzyme bound with N-(5′-phosphopyridoxyl)alanine . J Biol Chem 277 : 19166 – 19172 .

Watzele G Tanner W ( 1989 ) Cloning of the glutamine: fructose-6-phosphate amidotransferase gene from yeast. Pheromonal regulation of its transcription . J Biol Chem 264 : 8753 – 8758 .

Wild J Hennig J Łobocka M Walczak W Kłopotowski T ( 1985 ) Identification of the dadX gene coding for the predominant isozyme of alanine racemase in Escherichia coli K12 . Mol Gen Genet 198 : 315 – 322 .

Winterburn PJ Phelps CF ( 1971 ) The binding of substrates and modifiers to glucosamine synthetase . Biochem J 121 : 721 – 730 .

Wipat A Carter N Brignell SC Guy BJ Piper K Sanders J Emmerson PT Harwood CR ( 1996 ) The dnaB - pheA (256 degrees–240 degrees) region of the Bacillus subtilis chromosome containing genes responsible for stress responses, the utilization of plant cell walls and primary metabolism . Microbiology 142 : 3067 – 3078 .

Wong KK Kuo DW Chabin RM Fournier C Gegnas LD Waddell ST Marsilio F Leiting B Pompliano DL ( 1998 ) Engineering a cell-free murein biosynthetic pathway: combinatorial enzymology in drug discovery . J Am Chem Soc 120 : 13527 – 13528 .

Yamashita T Ashiuchi M Ohnishi K Kato S Nagata S Misono H ( 2003 ) Molecular characterization of alanine racemase from Bifidobacterium bifidum . J Mol Catal B Enzymatic 23 : 213 – 222 .

Yan Y Munshi S Leiting B Anderson MS Chrzas J Chen Z ( 2000 ) Crystal structure of Escherichia coli UDPMurNAc-tripeptide d-alanyl-d-alanine-adding enzyme (MurF) at 2.3 Å resolution . J Mol Biol 304 : 435 – 445 .

Yang Y Severin A Chopra R et al. ( 2006 ) 3,5-Dioxopyrazolidines, novel inhibitors of UDP- N -acetylenolpyruvylglucosamine reductase (MurB) with activity against Gram-positive bacteria . Antimicrob Agents Chemother 50 : 556 – 564 .

Yokoigawa K Hirasawa R Ueno H Okubo Y Umesako S Soda K ( 2001 ) Gene cloning and characterization of alanine racemases from Shigella dysenteriae , Shigella boydii , Shigella flexneri and Shigella sonnei . Biochem Biophys Res Commun 288 : 676 – 684 .

Yoshimura T Esaki N ( 2003 ) Amino acid racemases: functions and mechanisms . J Biosci Bioeng 96 : 103 – 109 .

Zawadzke LE Bugg TDH Walsh CT ( 1991 ) Existence of two d-alanine: d-alanine ligases in Escherichia coli : cloning and sequencing of the ddlA gene and purification and characterization of the DdlA and DdlB enzymes . Biochemistry 30 : 1673 – 1682 .

Zboińska E Sztajer H Lejczak B Kafarski P ( 1990 ) Antibacterial activity of phosphono dipeptides based on 1-amino-1-methylethanephosphonic acid . FEMS Microbiol Lett 58 : 23 – 28 .

Zeng B Wong KK Pompliano DL Reddy S Tanner ME ( 1998 ) A phosphinate inhibitor of the meso -diaminopimelic acid-adding enzyme (MurE) of peptidoglycan biosynthesis . J Org Chem 63 : 10081 – 10086 .

Zentz F Valla A Le Guillou R Labia R Mathot AG Sirot D ( 2002 ) Synthesis and antimicrobial activities of N-substituted imides . Farmaco 57 : 421 – 426 .

Ziegler K Diener A Herpin C Richter R Deutzmann R Lockau W ( 1998 ) Molecular characterization of cyanophycin synthetase, the enzyme catalyzing the biosynthesis of the cyanobacterial reserve material multi-l-arginyl-poly-l-aspartate (cyanophycin) . Eur J Biochem 254 : 154 – 159 .

Editor: Jacques Coyette

Email alerts

Citing articles via.

• Recommend to your Library
• Journals Career Network

Affiliations

• Online ISSN 1574-6976
• Print ISSN 0168-6445
• Copyright © 2024 Federation of European Microbiological Societies
• About Oxford Academic
• Publish journals with us
• University press partners
• What we publish
• New features  
• Open access
• Institutional account management
• Rights and permissions
• Get help with access
• Accessibility
• Advertising
• Media enquiries
• Oxford University Press
• Oxford Languages
• University of Oxford

Oxford University Press is a department of the University of Oxford. It furthers the University's objective of excellence in research, scholarship, and education by publishing worldwide

• Copyright © 2024 Oxford University Press
• Cookie settings
• Cookie policy
• Privacy policy
• Legal notice

This Feature Is Available To Subscribers Only

Sign In or Create an Account

This PDF is available to Subscribers Only

For full access to this pdf, sign in to an existing account, or purchase an annual subscription.

Cell Wall Synthesis

Cite this chapter.

• Angel Durán &
• Pilar Pérez  

462 Accesses

12 Citations

Cell growth in S. pombe is a complex process necessarily related to cell wall biosynthesis (Ishiguro 1998 , for review). The cell wall is the rigid structure that provides mechanical protection and supports the internal osmotic pressure, thereby restricting the entire morphology of this organism (Chap. 16). The cell wall must also be a dynamic structure to allow the morphological changes that occur during the life cycle, such as monopolar growth at first, then transition to bipolar growth upon ‘new end take-off’ (NETO), and cytokinesis by medial fission (Chap. 15), as well as mating of cells during zygote formation (Chap. 18) and spore wall formation around the postmeiotic nuclei, still surrounded by the outer ascus wall (Chap. 20).

This is a preview of subscription content, log in via an institution to check access.

Access this chapter

Subscribe and save.

• Get 10 units per month
• Download Article/Chapter or eBook
• 1 Unit = 1 Article or 1 Chapter
• Cancel anytime
• Available as PDF
• Read on any device
• Instant download
• Own it forever
• Compact, lightweight edition
• Dispatched in 3 to 5 business days
• Free shipping worldwide - see info
• Durable hardcover edition

Tax calculation will be finalised at checkout

Purchases are for personal use only

Institutional subscriptions

Unable to display preview.  Download preview PDF.

Similar content being viewed by others

Bacterial cell wall biogenesis is mediated by SEDS and PBP polymerase families functioning semi-autonomously

Stress-induced formation of cell wall-deficient cells in filamentous actinomycetes

Cell wall integrity signaling regulates cell wall-related gene expression in Chlamydomonas reinhardtii

Arellano M, Duran A, Pérez P (1996) Rhol GTPase activates the (1–3)/3-n-glucan synthase and is involved in Schizosaccharomyces pombe morphogenesis. EMBO J 15: 4584–4591

PubMed   CAS   Google Scholar  

Arellano M, Duran A, Pérez P (1997) Localization of the Schizosaccharomyces pombe rhol GTPase and its involvement in the organization of the actin cytoskeleton. J Cell Sci 110: 2547–2555

Arellano M, Coll PM, Pérez P (1999a) Rho GTPases in the control of cell morphology, cell polarity, and actin localization in fission yeast. Microsc Res Tech 47: 51–60

Article   PubMed   CAS   Google Scholar  

Arellano M, Valdivieso MH, Calonge TM et al. (1999 b) Schizosaccharomyces pombe protein kinase C homologues, pcklp and pcklp, are targets of rholp and rho2p and differentially regulate cell integrity. J Cell Sci 112: 3569–3578

Google Scholar  

Arellano M, Cartagena-Lirola H, Nasser Hajibagheri MA et al. (2000) Proper ascospore maturation requires the chsl’ chitin synthase gene in Schizosaccharomyces pombe . Mol Microbiol 35: 79–89

Ballou CE, Ballou L, Ball G (1994) Schizosaccharomyces pombe glycosylation mutant with altered cell surface properties. Proc Natl Acad Sci USA 91: 9327–9331

Banuett F (1998) Signalling in the yeast: an informational cascade with links to the filamentous fungi. Microbiol Mol Biol Rev 62: 249–274

Blad S, Kapteyn JC, Mol PC et al. (2001) Cell wall proteins of Schizosaccharomyces pombe . International Conference on Molecular mechanisms of fungal cell wall biogenesis: P17

Bowen AR, Chen-Wu JL, Momany M et al. (1992) Classification of fungal chitin synthases. Proc Natl Acad Sci USA. 89: 519–523

Bush DA, Horisberger M, Horman I, Wursch P (1974) The wall structure of S. pombe . J Gen Microbiol 81: 199–206

Cabib E, Drgonova J, Drgon T (1998) Role of small G proteins in yeast cell polarization and wall biosynthesis. Annu Rev Biochem 67: 307–333

Cabib E, Roh DH, Schmidt M et al. (2001) The yeast cell wall and septum as paradigms of cell growth and morphogenesis. J Biol Chem 276: 19679–19682

Calonge TM, Nakano K, Arellano M et al. (2000) Schizosaccharomyces pombe Rho2 GTPase regulates the cell wall a-glucan biosynthesis, through the protein kinase Pck2p. Mol Biol Cell 11: 4393–4401

Carnero E, Ribas JC, Garcia B et al. (2000) Schizosaccharomyces pombe ehslp is involved in maintaining cell wall integrity and in calcium uptake. Mol Gen Genet 264: 173–183

Castro C, Ribas JC, Valdivieso MH et al. (1995) Papulacandin B resistance in budding and fission yeasts: isolation and characterization of a gene involved in (1,3)ß-D-glucan synthesis in Saccharomyces cerevisiae. J. Bacteriol 177:5732–5739

Colussi PA, Orlean P (1997) The essential Schizosaccharomyces pombe gpil + gene complements a bakers’s yest GPI anchoring mutant and is required for efficient cell separation. Yeast 13:139–150

Cortés JC, Ishiguro J, Duran A, Ribas JC (2002) Location of the (1,3)/i-D-glucan synthase catalytic subunit homologue Bgslp/Cpslp from fission yeast suggests that it is involved in septation, polarized growth, mating, spore wall formation and spore germination. J Cell Sci 115: 4081–4096

Article   PubMed   Google Scholar  

Ding DQ, Tornita Y, Yamamoto A et al. (2000) Large-scale screening of intracellular protein localization in living fission yeast cells by the use of a GFP-fusion genomic DNA library. Genes Cells 5: 169–90

Feoktistova A, Magnelli P, Abeijon C et al. (2001) Coordination between fission yeast glucan

formation and growth requires a sphingolipase activity. Genetics 158:1397–1411

Gustin MC, Albertyn J, Alexander M, Davenport K (1998) MAP Kinase pathways in the Yeast Saccharomyces cerevisiae . Microbiol Mol Rev 62: 1264–1300

CAS   Google Scholar  

Heinisch JJ, Lorberg A, Schmitz HP, Jacoby JJ (1999) The protein kinase C-mediated MAP kinase pathway involved in the maintenance of cellular integrity in Saccharomyces cerevisiae. Mol Microbiol 32:671–680

Hirata D, Nakano K, Fukui M et al. (1998) Genes that cause aberrant cell morphology by overexpression in fission yeast: a role for a small GTP-binding protein Rho2 in cell morphogenesis. J Cell Sci 111: 149–159

Horisberger M, Rouvet-Vauthey M (1985) Cell wall architecture of the fission yeast Schizo-saccharomyces pombe. Experientia 41:748–749

Hochstenbach F, Klis FM, Van den Ende H et al. (1998) Identification of a putative alphaglucan synthase essential for cell wall construction and morphogenesis in fission yeast. Proc Natl Acad Sci USA 95: 9161–9166

Humbel BM, Konomi M, Takagi T et al. (2001) In situ localization of ß-glucans in the cell wall of Schizosaccharomyces pombe. Yeast 18:433 444

Ishiguro J (1998). Genetic control of fission yeast cell wall synthesis: the genes involved in wall biogenesis and their interactions in Schizosacharomyces pombe . Genes Genet Syst 73: 181–191

Ishiguro J, Saitou A, Duran A, Ribas JC (1997) cps1 + a Schizosaccharomyces pombe gene homolog of Saccharomyces cerevisiae FKS genes whose mutation confers hypersensitivity to cyclosporin A and papulacandin B. J Bacteriol 179:7653–7662

Ishijima SA, Konomi M, Takagi T et al. (1999) Ultrastructure of cell wall of the cps8 actin mutant cell in Schizosaccharomyces pombe. FEMS Microbiol Lett 180:31–37

Kanbe T, Kobayashi I, Tanaka K (1989) Dynamics of cytoplasmic organelles in the cell cycle of the fission yeast Schizosaccharomyces pombe : three-dimensional reconstruction from serial sections. J Cell Sci 94: 647–656

PubMed   Google Scholar  

Katayama S, Hirata D, Arellano M et al. (1999) Fission yeast)a-D-glucan synthase Mokl requires the actin cytoskeleton to localize the sites of growth and plays an essential role in cell morphogenesis downstream of protein kinase C function. J Cell Biol 144: 1173–1186

Kobori H, Yamaka N, Taki A, Osumi M (1989) Actin is associated with the formation of the cell wall in reverting protoplasts of the fission yeast Schizosaccharomyces pombe. J Cell Sci 94:635–646

Konomi M, Fujimoto K, Toda T, Osumi M (2001) a-1,3-Glucan is an essential component for cell wall formation of fission yeast. International Conference on Molecular Mechanisms of Fungal Cell Wall Biogenesis:P15

Kopecka M, Fleet GH, Phaff HJ (1995) Ultrastructure of the cell wall of Schizosaccharomyces pombe following treatment with various glucanases. J Struct Biol 114: 140–152

Le Goff X, Woollard A, Simanis V (1999) Analysis of the cpsl gene provides evidence for a septation checkpoint in Schizosaccharomyces pombe . Mol Gen Genet 262: 163–172

Lipke PN, Ovalle R (1998) Cell wall architecture in yeast: new structure and new challenges. J Bacteriol 180: 3735–3740

Liu J, Wang H, MacCollum D, Balasubramanian MK (1999) Drclp/Cpslp, a 1,3-ß-glucan synthase subunit, is essential for division septum assembly in Schizosaccharomyces pombe . Genetics 153: 1193–1203

Liu J, Tang X, Wang H, Balasubramanian M (2000 a) Bgs2p, a 1,3-ß-glucan synthase subunit, is essential for maturation of ascospore wall in Schizosaccharomyces pombe . FEBS Lett 478: 105–108

Liu J, Wang H, Balasubramanian MK (2000b) A checkpoint that monitors cytokinesis in Schizosaccharomyces pombe . J Cell Sci. 113: 1223–1230

Liu J, Tang X, Wang H et al. (2002) The location of the integral membrane protein Cpsl to the cell division site is dependent on the actomyosin ring and the septation inducing network in Schizosaccharomyces pombe . Mol Biol Cell 13: 989–1000

Manners DJ, Meyer MT (1977) The molecular structures of some glucans from the cell wall of S. pombe . Carbohyd Res 57: 189–203

Article   CAS   Google Scholar  

Martin V, Ribas JC, Carnero E et al. (2000) bgs2 + , a sporulation-specific glucan synthase homologue is required for proper ascospore wall maturation in fission yeast. Mol Microbiol 38: 308–321

Martin V, Garcia B, Carnero E et al. (2003) Bgs3p, a putative 1,313-glucan synthase subunit, is required for cell wall assembly in Schizosaccharomyces pombe . Eukar Cell 2:159–169 Martin-Cuadrado et al. (2003) Medline reference PMID: 12665550

Matsuo Y, Matasuda H, Kawamukai M (2001) Genetic analysis of chitin synthase I and II genes of fission yeast. Yeast 18: 5153.

Merla A, Johnson DI (2001) The Schizosaccharomyces pombe Cdc42p GTPase signals through Pak2p and the Mkhlp-Peklp-Spmlp MAPK pathway. Curr Genet 39: 205–209

Nakano K, Arai R, Mabuchi I (1997) The small GTP binding protein Rhol is a multifunctional protein that regulates actin localization, cell polarity, and septum formation in the fission yeast Schizosaccharomyces pombe . Genes Cells 2: 679–694

Nakano K, Mutoh T, Mabuchi I (2001) Characterization of GTPase-activating proteins for the function of the Rho-family small GTPases in the fission yeast Schizosaccharomyces pombe . Genes Cells 6: 1031–1042

Nurse P (1994) Fission yeast morphogenesis. Mol Biol Cell 5: 613–616

Osumi M (1998) The ultrastructure of yeast: cell wall structure and formation. Micron 29: 207–233.

Osumi M, Yamada N, Kobori H et al. (1989) Cell wall formation in regenerating protoplast of Schizosaccharomyces pombe : study by high resolution, low voltage scanning electron microscopy. J Electron Microsc 38: 457–468

Philip B, Levin DE (2001) Wscl and Mid2 are cell surface sensors for cell wall integrity signaling that act through Rom2, a guanine nucleotide exchange factor for Rhol. Mol Cell Biol 21: 271–280

Popolo L, Vai M (1999) The Gasl glycoprotein, a putative wall polymer cross-linker. Biochim Biophys Acta 1426: 385–400

Ram AF, Kapteyn JC, Montijn RC et al. (1998) Loss of the plasma membrane-bound protein Gaslp in Saccharomyces cerevisiae results in the release of betal,3-glucan into the medium and induces a compensation mechanism to ensure cell wall integrity. J Bacteriol 180: 1418–1824

Ribas JC, Díaz M, Duran A, Pérez P (1991) Isolation and characterization of Schizosaccharo- myces pombe mutants defective in cell wall (1–3)ß-D-glucan. J Bacteriol 173: 3456–3462

Robinow CF, Hyams JS (1989) General cytology of fission yeast. In: Nasim A, Young P, Johnson BF (eds) Molecular biology of the fission yeast. Academic Press, London, pp 273330

Sayers LG, Katayama S, Nakano K et al. (2000) Rho-dependence of Schizosaccharomyces pombe Pck2. Genes Cells 5: 17–27

Sells MA, Barratt JT, Caviston J et al. (1998) Characterization of Pak2p, a pleckstrin homology domain-containing, p21-activated protein kinase from fission yeast. J Biol Chem 273: 18490–18498

Sengar AS, Markley NA, Marini NJ, Young D (1997) Mkhl, a MEK kinase required for cell wall integrity and proper response to osmotic and temperature stress in Schizosaccharomyces pombe . Mol Cell Biol 17: 3508–3519

Shiozaki K, Russell P (1995) Counteractive roles of protein phosphatase 2C (PP2C) and a

MAP kinase kinase homolog in the osmoregulation of fission yeast. EMBO J 14:492–502 Sietsma JH, Wessels JGH (1990) The occurrence of glucosaminoglycan in the wall of S. pombe . J Gen Microbiol 136: 2261–2265

Sugawara T, Takagi T, Sato M et al. (2001) Architecture and localization of the cell wall glu-cans of the fission yeast, Schizosaccharomyces pombe . International Conference on Molecular mechanisms of fungal cell wall biogenesis: P16

Sugiura R, Toda T, Dhut S et al. (1999) The MAPK kinase Pekl acts as a phosphorylationdependent molecular switch. Nature 399: 479–483

Toda T, Shimanuki M, Yanagida M (1993) Two novel protein kinase C-related genes of fission yeast are essential for cell viability and implicated in cell shape control. EMBO J 12: 1987–1995

Toda T, Dhut S, Superti-Furga G et al. (1996) The fission yeast pmk1 + gene encodes a novel mitogen-activated protein kinase homolog which regulates cell integrity and functions coordinately with the protein kinase C pathway. Mol Cell Biol 16: 6752–6764

Varona R, Pérez P, Duran A (1983) Effect of papulacandin B on /3-glucan synthesis in Schizosaccharomyces pombe . FEMS Microbiol Lett 20: 243–247

Yang P, Kansra S, Pimental RA et al. (1998) Cloning and characterization of shk2 , a gene encoding a novel p21-activated protein kinase from fission yeast. J Biol Chem 273: 1848118489

Zaitsevskaya-Carter T, Cooper JA (1997) Spml, a stress-activated MAP kinase that regulates morphogenesis in S. pombe . EMBO J 16: 1318–1331

Download references

You can also search for this author in PubMed   Google Scholar

Editor information

Editors and affiliations.

Department of Genetics, Institute of Molecular Biology, University of Copenhagen, Øster Farimagsgade 2A, 1353, Copenhagen K, Denmark

Richard Egel

Rights and permissions

Reprints and permissions

Copyright information

© 2004 Springer-Verlag Berlin Heidelberg

About this chapter

Durán, A., Pérez, P. (2004). Cell Wall Synthesis. In: Egel, R. (eds) The Molecular Biology of Schizosaccharomyces pombe . Springer, Berlin, Heidelberg. https://doi.org/10.1007/978-3-662-10360-9_17

Download citation

DOI : https://doi.org/10.1007/978-3-662-10360-9_17

Publisher Name : Springer, Berlin, Heidelberg

Print ISBN : 978-3-642-05631-4

Online ISBN : 978-3-662-10360-9

eBook Packages : Springer Book Archive

Share this chapter

Anyone you share the following link with will be able to read this content:

Sorry, a shareable link is not currently available for this article.

Provided by the Springer Nature SharedIt content-sharing initiative

• Publish with us

Policies and ethics

• Find a journal
• Track your research

• Target Discovery for Cancer Immunotherapy
• Identification of Biomarkers by ORFeome Phage Display
• Whole Genome Sequencing (WGS)
• In Silico Target Prediction
• In Silico Protein Modeling
• In Silico Functional Analysis for Fungi
• Gene Expression Profiling
• Affinity Chromatography
• Gene Knockout
• Gene Conditional Expression
• Gene Overexpression Service
• X-ray Crystallography
• Nuclear Magnetic Resonance (NMR) Spectroscopy
• Cryo-Electron Microscopy
• Small-Angle X-ray and Neutron Scattering
• Digital Imaging
• Metabolic Pathway Analysis for Fungi
• Target Localization
• In Vivo Target Validation
• Cell Wall Targets Identification
• Virtual Screening
• Compound Libraries
• Antibody Discovery
• High Throughput Screening
• Fragment-Based Screening
• Phenotypic Screening
• Secondary Screening
• Drug Repurposing
• Hit Characterization
• Developability Assessment of Biopharmaceutical Candidates
• Molecular Engineering
• Antibody Expression
• In Vitro Pharmacodynamics
• Anti-Drug Antibody (ADA) Services
• In Vitro Safety Pharmacology Studies
• In Vitro ADME
• In Vitro Toxicity Test
• In Vivo PK Studies
• Pharmacology and Pharmacodynamics Studies
• Animal Models Service
• Drug Abuse Liability
• Pre-formulation Development
• Formulation Development
• Human Dose Prediction
• Process and Scale-up
• Complement Therapeutic Services
• Antifungal Drug Discovery
• Antibacterial Drug Discovery
• MIB-CM ® Dry Powder Medium
• MIB-CM ® Chromogenic Medium
• Function Assay Kits
• Function Assay

• Hit to Lead
• Antibacterial MOA Determination

Interference with Cell Wall Synthesis

The knowledge of the mode of action (MOA) is crucial for discovering new antibacterial drugs. The MOA of antibiotics explains how antibiotics can attack bacteria without affecting human cells. Creative Biolabs has developed a top drug discovery platform to determine the mode of action of antibacterial drugs in cell wall synthesis.

Bacterial cells are different from animal cells as they have cell walls. Therefore, a drug that attacks cell walls will have no significant negative effect on the cells of the mammalian host. The cell wall is composed of units of peptides (proteins) and glycans (sugars). Most bacteria have cell walls consisting of single or more protective layers on top of their cytoplasmic membranes. Gram-negative bacteria also have an additional outer lipid bilayer. Cell wall protects bacteria from injury. Inhibit cell wall synthesis is one of the most common modes of action for antibiotics. By inhibiting peptidoglycan synthesis, the growth of bacteria is prevented. These bacteria will be subjected to osmotic lysis.

Generally, the modes of actions of antibiotics for interference with cell wall are the following:

Interference with Peptidoglycan

Peptidoglycan is a promising target in bacterial cell wall for broad-spectrum antibacterial drug discovery, as the disaccharide-pentapeptide peptidoglycan structure is common to both Gram-negative and Gram-positive bacteria. Many antibiotics have effects on different stages of peptidoglycan, either synthesis or transport of its metabolic precursors or by direct action on its structural organization. For instance, beta-lactams Inhibit synthesis of peptidoglycan; vancomycin disrupts peptidoglycan cross-linkage; bacitracin disrupts the movement of peptidoglycan precursors. Knowing the mode of action of new antibiotics, we may also have some knowledge of mechanisms of resistance.

Interference with Teichoic Acids (TA)

Except for peptidoglycan, Gram-positive bacteria are comprised of a second, equally abundant component: teichoic acid. Teichoic acid is comprised of linear, polyol phosphate polymers that exist in one of two forms: wall teichoic acid (WTA) attached to wall peptidoglycan, and lipoteichoic acid (LTA) linked to membrane lipids. WTA are involved in many aspects of cell division and are essential for maintaining cell shape in rod-shaped organisms. LTA plays an important role in bacterial growth and physiology and contributes to membrane homeostasis. Both WTA and LTA are potential targets for new therapeutics to overcome bacterial infections.

Interference with Lipopolysaccharides (LPS)

Lipopolysaccharides (LPS) (also lipoglycans or endotoxins) are a kind of large molecules consisting of a lipid and a polysaccharide. They are important components in the outer membrane of Gram-negative bacteria and can act as antigens that elicit strong immune responses in animals. Both LPS synthesis and LPS transport are of crucial importance to Gram-negative bacteria. Interference of LPS is also an important MOA for antibiotics.

Creative Biolabs is a trusted partner in identifying the mode of action for novel antibiotics. For more detailed information, please feel free to contact us or directly sent us an inquiry.

For Research Use Only.

• Remote Access
• Save figures into PowerPoint
• Download tables as PDFs

Chapter 28:  Inhibitors of Cell Wall Synthesis

• Download Chapter PDF

Disclaimer: These citations have been automatically generated based on the information we have and it may not be 100% accurate. Please consult the latest official manual style if you have any questions regarding the format accuracy.

Download citation file:

• Search Book

Jump to a Section

General features.

• OTHER INHIBITORS OF CELL WALL SYNTHESIS
• Full Chapter
• Supplementary Content

These drugs all work by inhibiting the synthesis of the bacterial cell wall.

You probably guessed this from the title of the chapter. However, this is a real key point. If you can remember this, you are well on your way to learning these drugs.

The final step in the synthesis of the bacterial cell wall is a cross-linking of adjacent peptidoglycan strands by a process called transpeptidation . The penicillins and cephalosporins are structurally similar to the terminal portion of the peptidoglycan strands and can compete for and bind to the enzymes that catalyze transpeptidation and cross-linking. These enzymes are called penicillin-binding proteins (PBPs). Interference with these enzymes results in the formation of a structurally weakened cell wall, oddly shaped bacteria, and ultimately, death.

Now, let’s divide the cell wall synthesis inhibitors into two groups based on chemical structure: β-lactams and others.

β-Lactam antibiotics are best against rapidly growing organisms. They are not effective against intracellular organisms.

This is where your knowledge of the microbes can really help you. β-Lactam antibiotics interfere with the synthesis of the cell wall, which is on the outside of the bacteria. Rapidly growing bacteria are making lots of cell walls making them susceptible to these antibiotics.

All of the drugs in this group contain a β-lactam ring in their structure.

Normally, we do not worry too much about the structures of drugs, but in this case we make an important exception. These drugs are often referred to as the β-lactam group. This is because they all have a β-lactam ring in their chemical structure, and it is this β-lactam ring that makes them effective antimicrobials.

Some bacteria inactivate β-lactam antibiotics by an enzyme that opens the β-lactam ring.

Some bacteria contain an enzyme, called β-lactamase , that can open the β-lactam ring ( Figure 28–1 ). This leads to inactivation of the antibiotic. The most common mode of drug resistance is plasmid transfer of the genetic code for the β-lactamase enzyme. There is a β-lactamase specific for the penicillins—it is called penicillinase —and a β-lactamase specific for the cephalosporins—it is called cephalosporinase . Inactivation of these drugs by β-lactamases is a major problem and has been the focus of intense research.

FIGURE 28–1

Here we can see the lactam ring and its opening by penicillinase.

The inactivation of these drugs by β-lactamases can be dealt with by two approaches:

Give a β-lactamase inhibitor at the same time.

Make chemical modifications in the structure of the drug to make it more resistant to inactivation.

CLAVULANIC ACID and SULBACTAM are β-lactamase inhibitors that are given together with β-lactam drugs to increase their effectiveness.

Get Free Access Through Your Institution

Pop-up div successfully displayed.

This div only appears when the trigger link is hovered over. Otherwise it is hidden from view.

Please Wait

An official website of the United States government

The .gov means it’s official. Federal government websites often end in .gov or .mil. Before sharing sensitive information, make sure you’re on a federal government site.

The site is secure. The https:// ensures that you are connecting to the official website and that any information you provide is encrypted and transmitted securely.

• Publications
• Account settings
• My Bibliography
• Collections
• Citation manager

Save citation to file

Email citation, add to collections.

• Create a new collection
• Add to an existing collection

Add to My Bibliography

Your saved search, create a file for external citation management software, your rss feed.

• Search in PubMed
• Search in NLM Catalog
• Add to Search

Inhibition of cell wall synthesis--is this the mechanism of action of penicillins?

Affiliation.

• 1 Medical Division, Unichem Laboratories Ltd., Jogeshwari, (West) Bombay, India.
• PMID: 7596307
• DOI: 10.1016/0306-9877(95)90085-3

Penicillins have been shown to inhibit bacterial cell wall synthesis, and interact with penicillin binding proteins, leading to bacterial lysis. These two mechanisms, the former more than the latter are believed to be responsible for their therapeutic potential. It has further been demonstrated that only actively multiplying cells are susceptible to bactericidal effects of the antibiotic, which is in accordance with the suggested mechanism of action. Bacterial growth takes place in terms of size and number, both requiring additional cell wall. An increase in bacterial size is due to an increase in the volume of cytosol and area of the cell wall. Presently there is no proof that the former is the cause of the latter or vice versa. Penicillin by inhibiting cell wall synthesis would inhibit both growth and multiplication. Since the antibiotic is bactericidal to rapidly multiplying cells, its effect on cell wall would interfere with its bactericidal action. As per the present understanding penicillin acts principally by inhibiting cell wall synthesis. There is however a discrepancy between its observed effects and what should logically be expected, which forces us to reexamine the mechanism of action of penicillin. We believe that the present understanding of the action of penicillin is incomplete if not outright faulty. It would be expedient to radically modify the same in view of its implication, for example on drug development.

PubMed Disclaimer

Similar articles

• An experimental analysis of the curative action of penicillin in acute bacterial infections. III. The effect of suppuration upon the antibacterial action of the drug. SMITH MR, WOOD WB Jr. SMITH MR, et al. J Exp Med. 1956 Apr 1;103(4):509-22. doi: 10.1084/jem.103.4.509. J Exp Med. 1956. PMID: 13306859 Free PMC article.
• Defective killing of enterococci: a common property of antimicrobial agents acting on the cell wall. Krogstad DJ, Pargwette AR. Krogstad DJ, et al. Antimicrob Agents Chemother. 1980 Jun;17(6):965-8. doi: 10.1128/AAC.17.6.965. Antimicrob Agents Chemother. 1980. PMID: 6902640 Free PMC article.
• Mechanism of action of penicillin: triggering of the pneumococcal autolytic enzyme by inhibitors of cell wall synthesis. Tomasz A, Waks S. Tomasz A, et al. Proc Natl Acad Sci U S A. 1975 Oct;72(10):4162-6. doi: 10.1073/pnas.72.10.4162. Proc Natl Acad Sci U S A. 1975. PMID: 674 Free PMC article.
• Penicillin-binding proteins and role of amdinocillin in causing bacterial cell death. Neu HC. Neu HC. Am J Med. 1983 Aug 29;75(2A):9-20. doi: 10.1016/0002-9343(83)90089-x. Am J Med. 1983. PMID: 6311012 Review.
• [Penetration, site and mode of action of antibiotics in bacteria]. Videau D. Videau D. Biol Med (Paris). 1968 Nov-Dec;57(6):449-70. Biol Med (Paris). 1968. PMID: 4238452 Review. French. No abstract available.
• The Benefits and Challenges of Antibiotics-Non-Steroidal Anti-Inflammatory Drugs Non-Covalent Reaction. Nugrahani I, Herawati D, Wibowo MS. Nugrahani I, et al. Molecules. 2023 Apr 23;28(9):3672. doi: 10.3390/molecules28093672. Molecules. 2023. PMID: 37175082 Free PMC article. Review.
• Hylin-a1: A Host Defense Peptide with Antibacterial Potential against Staphylococcus aureus Multi-Resistant Strains. Chianese A, Zannella C, Foglia F, Nastri BM, Monti A, Doti N, Franci G, De Filippis A, Galdiero M. Chianese A, et al. Pharmaceuticals (Basel). 2023 Mar 29;16(4):509. doi: 10.3390/ph16040509. Pharmaceuticals (Basel). 2023. PMID: 37111266 Free PMC article.
• An antibacterial compound pyrimidomycin produced by Streptomyces sp. PSAA01 isolated from soil of Eastern Himalayan foothill. Das P, Kundu S, Maiti PK, Mandal S, Sahoo P, Mandal S. Das P, et al. Sci Rep. 2022 Jun 17;12(1):10176. doi: 10.1038/s41598-022-14549-4. Sci Rep. 2022. PMID: 35715695 Free PMC article.
• Bis-Indole Alkaloids Isolated from the Sponge Spongosorites calcicola Disrupt Cell Membranes of MRSA. Khan NA, Kaur N, Owens P, Thomas OP, Boyd A. Khan NA, et al. Int J Mol Sci. 2022 Feb 11;23(4):1991. doi: 10.3390/ijms23041991. Int J Mol Sci. 2022. PMID: 35216106 Free PMC article.
• Smarter cures to combat COVID-19 and future pathogens: a review. Dai H, Han J, Lichtfouse E. Dai H, et al. Environ Chem Lett. 2021;19(4):2759-2771. doi: 10.1007/s10311-021-01224-9. Epub 2021 Apr 2. Environ Chem Lett. 2021. PMID: 33824633 Free PMC article. Review.
• Search in MeSH

Related information

• Cited in Books
• PubChem Compound (MeSH Keyword)

LinkOut - more resources

Full text sources.

• Elsevier Science

Research Materials

• NCI CPTC Antibody Characterization Program
• Citation Manager

NCBI Literature Resources

MeSH PMC Bookshelf Disclaimer

The PubMed wordmark and PubMed logo are registered trademarks of the U.S. Department of Health and Human Services (HHS). Unauthorized use of these marks is strictly prohibited.

An official website of the United States government

The .gov means it’s official. Federal government websites often end in .gov or .mil. Before sharing sensitive information, make sure you’re on a federal government site.

The site is secure. The https:// ensures that you are connecting to the official website and that any information you provide is encrypted and transmitted securely.

• Publications
• Account settings

Preview improvements coming to the PMC website in October 2024. Learn More or Try it out now .

• Advanced Search
• Journal List
• Arabidopsis Book

Cellulose Synthesis and Its Regulation

a Department of Biochemistry and Molecular Biology, The Pennsylvania State University, University Park, PA 16802

Logan Bashline

Cellulose, the most abundant biopolymer synthesized on land, is made of linear chains of ß (1–4) linked D-glucose. As a major structural component of the cell wall, cellulose is important not only for industrial use but also for plant growth and development. Cellulose microfibrils are tethered by other cell wall polysaccharides such as hemicellulose, pectin, and lignin. In higher plants, cellulose is synthesized by plasma membrane-localized rosette cellulose synthase complexes. Despite the recent advances using a combination of molecular genetics, live cell imaging, and spectroscopic tools, many aspects of the cellulose synthesis remain a mystery. In this chapter, we highlight recent research progress towards understanding the mechanism of cellulose synthesis in Arabidopsis.

1. INTRODUCTION

Every cell in higher plants is encased in an extracellular matrix, the plant cell wall. The plant cell wall is composed of a mixture of polysaccharides ( Cosgrove, 2005 ). Based on structural and functional differences, plant cell walls can be roughly characterized into two types: the primary cell wall and the secondary cell wall. The composition and structural organization of both primary and secondary cell walls has been a major driving force for the evolutionary diversification of plants on earth ( Popper, 2008 ). The primary cell wall contains cellulose, hemicellulose and pectin. Primary cell wall synthesis occurs during the growing phase of the cell when the cell wall expands due to the forces of internal turgor pressure that push outward against the plasma membrane and the cell wall ( Cosgrove, 1997 ). The secondary cell wall is deposited in specific cell types upon the cessation of cell growth. The secondary cell wall contains cellulose, hemicellulose, and lignin and it is usually thicker and more rigid than the primary cell wall ( Roberts et al., 2000 ).

Through intra- and inter-chain hydrogen bonding, parallel linear glucan chains are crystalized to form cellulose microfibrils, which give cellulose high axial stiffness ( Gillis et al., 1969 ). In higher plants, 3-nm elementary cellulose microfibrils can aggregate into larger size cellulose microfibrils that range from 5–10 nm in width in primary cell walls to 30–50 nm in secondary cell walls ( Davies et al., 2003 ; Zhang, 2013 ) ( Figure 1 ). The length of cellulose microfibrils, which can be referred to as the degree of polymerization (DP), varies significantly among organisms. The DP of cellulose microfibrils is estimated to range from hundreds to thousands of glucose units in primary walls, and up to 15,000 glucose units in secondary walls ( Brett, 2000 ; Somerville, 2006 ). However, virtually nothing is known about how the DP is regulated in plants.

Cellulose microfibrils and cellulose synthase complexes (CSCs) in Arabidopsis. (A) Atomic force micrograph of cellulose microfibrils from Arabidopsis callus tissue after cyclohexane-1,2-diaminetetraacetic acid (CDTA) extraction. Image size is 2×2 µm. Adapted from Davies LM & Harris PJ ( 2003 ). Copyright by Springer. (B) Electron micrograph of cellulose microfibrils in Arabidopsis. Scale bar = 100 nm. Layers of cellulose microfibrils organized as sheets can be seen. Adapted from Williamson et al. ( 2002 ). Copyright by Elsevier. (C) Cellulose synthase complexes (CSCs) in the plasma membrane of Arabidopsis. Scale bar =100 nm. Adapted from Williamson et al. ( 2002 ). Copyright by Elsevier.

All cellulose-synthesizing organisms including bacteria, algae, tunicates, and higher plants have cellulose synthase proteins, which catalyze the polymerization of glucan chains ( Brett, 2000 ; Saxena et al., 2005 ). Although the catalytic domains of cellulose synthases are conserved for all cellulose-synthesizing organisms, the drastic differences in both the lifestyle of the organisms and the structure of the cellulose that they produce suggest that the regulatory proteins and the underlying mechanisms for cellulose synthesis may have evolved independently ( Roberts et al., 2007 ; Lei et al., 2012b ). An example of diversity is the variation in the organization of cellulose synthesizing complexes, which were originally named terminal complexes (TCs) due to their association with the ends of cellulose microfibrils ( Montezinos et al., 1976 ). TCs in higher plants adopt a rosette shape with six lobes that have rotational symmetry and span across the plasma membrane with a 25 nm diameter in the transmembrane region and a larger diameter (45–50 nm) in the cytoplasmic region of the complex ( Giddings et al., 1980 ; Mueller et al., 1980 ; Brown et al., 1996 ; Bowling et al., 2008 ). The protein composition of the rosette in higher plants is not well understood, but each rosette contains multiple cellulose synthase proteins to accommodate the synthesis of multiple glucan chains ( Taylor et al., 2000 ; Doblin et al., 2002 ; Desprez et al., 2007 ; Persson et al., 2007 ). It has been postulated that cellulose synthase complexes are involved not only in the polymerization of glucan chains, but also in the crystallization process ( Somerville, 2006 ). It remains unclear how multiple glucan chains are positioned within proximity of one another to accommodate crystallization through hydrogen bonding. It has been proposed that protein other than cellulose synthases may be a part of the complex and aid in the crystallization process in Arabidopsis ( Molhoj et al., 2002 ).

Cellulose biosynthesis in higher plants is a tightly regulated process ( Brabham et al., 2012 ; Li et al., 2012b ). For example, the amount of cellulose, the ratio of cellulose to other cell wall polymers, the degree of polymerization, the crystalline cellulose core size, and the orientation of cellulose microfibrils are under tight control. The vast regulation of cellulose biosynthesis is a reflection of the variability in cell wall structure and composition that arises from a diversity of cell types across various developmental stages ( Keegstra, 2010 ). This area has been greatly advanced by genetic studies in Arabidopsis and by the recent development of an in vivo cell imaging system to visualize the dynamics of cellulose synthase complexes (CSCs) in living cells. However, at the biochemical level, the field is still at the beginning stage of understanding the enzymatic mechanism of cellulose synthesis. In that regard, it is appropriate to present information from other species, even distant ones, to use as an introduction, comparison or reference to understand cellulose biosynthesis in Arabidopsis.

2. THE STRUCTURE AND PROPERTIES OF CELLULOSE

2.1. general structure of cellulose.

Cellulose, in its simplest form, consists of β-1,4 linked glucan chains. Hydrogen bonds form between hydroxyl groups and oxygen atoms both within a single glucose chain and between neighboring chains. Together with van der Waals forces, hydrogen bonding aggregates glucan chains together side-by-side and promotes parallel stacking of cellulose microfibrils into crystalline cellulose ( Brett, 2000 ; Somerville, 2006 ). The natural form of crystalline cellulose is cellulose I. Cellulose I can be irreversibly converted into cellulose II, a form that is more stable than the cellulose I ( Brown, 1999a ; Brett, 2000 ). Consisting of cellulose la and Iβ, both forms are composed of parallel glucan chains ( Kuga et al., 1988 ; Maurer et al., 1992 ; Koyama et al., 1997 ). The parallel glucan chains in natural cellulose are compatible with the idea that glucan chains in an elementary cellulose microfibril are made simultaneously ( Brown, 1999a ; Brown et al., 2000 ). In higher plants, the ratio of cellulose la to Iβ varies among different species and types of walls ( Atalla et al., 1984 ; Sturcova et al., 2004 ). In both forms, each glucose molecule is rotated 180° in relation to its neighboring glucose molecule, forming a flat ribbon in which cellobiose is the repeating unit ( Somerville, 2006 ).

Multiple glucan chains are simultaneously synthesized by a single plasma membrane-localized CSC and immediately assemble to form an elementary cellulose microfibril ( Harris et al., 2010 ). The size of an elementary fibril is physically determined by the number of individual glucan chains synthesized by a CSC ( Tsekos, 1999 ). In Arabidopsis, CSCs exist as rosettes ( Figure 1 ), which contain six subunits arranged in a hexagonal structure. It has been postulated that each of the six rosette subunits contains six cellulose synthase (CESA) proteins ( Doblin et al., 2002 ). Assuming that each CESA protein within a CSC is enzymatically active, this model would suggest that each CSC synthesizes an elementary microfibril that is comprised of 36 glucan chains ( Somerville, 2006 ). The exact number of individual glucan chains in an elementary cellulose microfibril has not been experimentally determined in Arabidopsis. However, studies on cellulose in celery collenchyma (primary wall) and in spruce wood (secondary wall) suggest that an elementary microfibril is more likely to contain 24 glucan chains ( Fernandes et al., 2011 ; Newman et al., 2013 ; Thomas et al., 2013 ). It is not clear whether the number of chains in elementary microfibrils is fixed within an organism let alone across species, but based on the current measurements of elementary cellulose microfibril size, it is reasonable to estimate that each of the six subunits of the CSC contains 4–6 enzymatically active CESAs.

Several methods have been developed to measure the crystallinity of cellulose microfibrils. X-ray diffraction (XRD) can provide a rough estimation of crystallinity through measuring the relative crystallinity index, which is based upon the proportions of crystalline and amorphous cell wall material, a measurement that can be influenced by non-cellulosic polysaccharides ( L. Segal, 1959 ; Harris et al., 2010 ; Park et al., 2010 ; Harris et al., 2012 ). Cellulose crystallinity can also be assessed by 13 C solid-state NMR spectroscopy, by comparing the relative intensities of peaks that correspond to C4 atoms in the interior of the cellulose versus C4 atoms that are on the surface of the cellulose microfibril, which can be used to estimate crystallite size ( Bootten et al., 2011 ; Dick-Perez et al., 2011 ). However, this method has trouble differentiating between amorphous cellulose regions and cellulose chains at the surface of crystalline microfibrils. 13 C solid-state NMR spectroscopy is best suited for crystallinity analysis of thick cellulose microfibrils (10–25 nm in diameter) from bacteria and certain algae ( Brett, 2000 ). However, cellulose microfibrils from the primary cell walls in higher plants are relatively thin, ranging from 3–10 nm in diameter ( Thomas et al., 2013 ; Zhang, 2013 ). A model in which regions of crystalline cellulose are interconnected by amorphous cellulose regions has gained popularity in recent years. These amorphous regions can be detected by small-angle neutron scattering and are highly accessible through acid hydrolysis ( Fernandes et al., 2011 ). However, it is important to note that the isolation of cellulose microfibrils in these studies often involves harsh extraction methods that will affect the native eellulose structure ( Somerville, 2006 ). A sum frequency generation (SFG) spectroscopy was recently used to detect the asymmetric distribution of C6H2 and O3H-O5 group in crystalline cellulose microfibrils ( Hieu et al., 2011 ; Barnette et al., 2012 ). SFG is a desirable method to estimate the content of crystalline cellulose because there is no spectral interference from other cell wall matrix compounds such as hemicellulose and lignin and it does not require any chemical treatment of biological samples ( Barnette et al., 2012 ). SFG can also detect subtle changes in cellulose ordering and packing in secondary cell wall in Arabidopsis inflorescence stem ( Park et al., 2013 ).

2.2. Cellulose microfibril organization

In general, cellulose microfibrils are laid down transversely to the axis of elongation during primary cell wall synthesis ( Figure 1 ). In Arabidopsis, two experimental systems, the epidermal cells of dark grown hypocotyls and the root elongation zone, have been used to investigate the relationship between cellulose biosynthesis and cell elongation of primary cell walls ( Fagard et al., 2000a ; Williamson et al., 2002 ). Rapid longitudinal cell elongation rates coupled with minimal lateral expansion make both systems ideal for genetic, physiological and structural analysis ( Crowell et al., 2010a ). In epidermal cells of the root elongation zone, newly synthesized cellulose is constantly deposited in a transverse orientation to the root axis ( Kerstens et al., 2002 ). During cell elongation, layers of cellulose microfibrils rotate as sheets with angles that increasingly move towards the longitudinal axis ( Anderson et al., 2010 ). The rotation of cellulose microfibril layers in roots is compatible with the multi-net growth hypothesis in which the growth is associated with passive movement of cell wall layers ( Roland et al., 1975 ). Cellulose microfibrils are also arranged in multilayer sheets of varying angles during hypocotyl elongation. In this cell type, cellulose microfibrils in the outer face of the epidermal cell wall are deposited in parallel with the underlying cortical microtubules that undergo continuous rotation ( Chan et al., 2007 ; Lloyd et al., 2008 ; Chan et al., 2010 ). It appears that these two systems have co-evolved to adopt a multi-net cellulose microfibril network. How multi-net cellulose microfibril organization relates to the features of the primary cell walls, especially extensibility and rigidity, are not known. Interestingly, secondary cell walls of wood cells also adopt a similar cellulose microfibril organization. During wood formation, cellulose microfibrils are laid down as three consecutive layers (S1, S2, S3). Within each layer the cellulose microfibrils are highly ordered and parallel, but between each layer the angles of the microfibril sheets are oriented differently ( Plomion et al., 2001 ; Barnett et al., 2004 ). Along with embedded hemicellulose (xylan) and ligin, the multi-layered cellulose microfibril organization makes the wood cell wall an ideal structure for strength and rigidity ( Chaffey, 1999 ; Plomion et al., 2001 ). Multilayer cellulose microfibril organization may be a convergent evolutionary product that is best fit for providing rigidity to the cell wall.

In both primary and secondary cell walls, cellulose microfibrils often exist as bundles ( Anderson et al., 2010 ; Fernandes et al., 2011 ; Thomas et al., 2013 ; Zhang, 2013 ). The bundling process likely involves the aggregation of closely arranged cellulose microfibrils. Since the geometry and dimensions of CSCs relate to the size of cellulose microfibrils, it is tempting to speculate that the bundling process might also be influenced by how the CSCs are arranged at the plasma membrane. Since cortical microtubules guide the insertion and movement of CSCs at the plasma membrane ( Crowell et al., 2009 ; Gutierrez et al., 2009 ), it is possible that as multiple CSCs simultaneously extrude cellulose microfibrils along the same cortical microtubule, these cellulose microfibrils interact with one another to form a larger and bundled cellulose microfibril. Therefore, it is reasonable to speculate that the dynamics of cortical microtubules might influence the bundling process. It is not known if the bundling process occurs simultaneously during individual glucan-chain synthesis or occurs sequentially after the termination of synthesis.

Other cell wall polymers that are in close contact with cellulose microfibrils such as hemicellulose and pectin may also affect the bundling of cellulose microfibrils. Xyloglucan is the most dominant hemicellulose in dicot primary walls and it plays a large role in forming the cross-links between cellulose microfibrils ( Keegstra et al., 1973 ). Cellulose microfibrils are also coated with xylogalacturonan, a pectin polysaccharide, during the synthesis of cellulose microfibrils in quince seed mucilage. The xylogalacturonan coating prevents the coalescence of cellulose microfibrils into large bundles ( Ha et al., 1998 ). In primary cell walls of Arabidopsis, pectin directly interacts with cellulose microfibrils. It has been estimated that 25–50% of the cellulose chains are in close contact with pectin polysaccharides ( Wang et al., 2012 ). While numerous documented studies support the direct interaction between hemicellulose/pectin and cellulose microfibrils ( Keegstra et al., 1973 ; Cosgrove, 1997 ; Keegstra, 2010 ; Scheller et al., 2010 ), it is not known how or to what extent the direct contact between these cell wall polymers affects the bundling process.

3. CELLULOSE SYNTHESIS

3.1. cesa genes, cesa proteins and the cellulose synthase complex (csc).

In 1996, the first higher plant cellulose synthase gene was cloned from cotton ( Pear et al., 1996 ). Since then, CESA genes have been identified in many higher plants including a few model systems such as Arabidopsis, rice, and poplar ( Carroll et al., 2011 ). CESA proteins of higher plants share relatively low amino acid sequence identity with bacterial CESA proteins ( Doblin et al., 2002 ), in which the CESA was first identified. In bacteria, the CESA gene is in an operon that also contains other genes that are directly related to cellulose synthesis and cellulose microfibril assembly ( Ross et al., 1991 ). However, in higher plants no functionally linked genes are present in the proximity of the CESA genes. CESAs belong to family 2 glycosyltransferases ( Richmond et al., 2000 ; Somerville, 2006 ). The predicted CESA protein structure suggests that some aspects of CESA structure are conserved between prokaryotes and eukaryotes ( Lei et al., 2012b ; Morgan et al., 2013 ; Sethaphong et al., 2013 ). CESAs are integral plasma membrane proteins with multiple transmembrane domains and a central catalytic domain ( Richmond, 2000 ) ( Figure 2 ). In higher plants, CESAs have eight transmembrane domains: two on the N-terminal side of the central domain and six on the C terminal side ( Somerville, 2006 ). These transmembrane domains are predicted to form a pore in the plasma membrane, which might accommodate the extrusion of newly synthesized glucan chains through the membrane ( Morgan et al., 2013 ; Slabaugh et al., 2013 ). In higher plants, CESAs contain a cytoplasmic amino terminal zinc finger domain, which is believed to play a role in protein-protein interactions and might be responsible for the dimerization of CESA proteins ( Kurek et al., 2002 ). Following the zinc finger domain is a hyper-variable region that is rich in acidic amino acids. In between the second and third transmembrane domains is the central domain or the globular domain, which extends into the cytoplasm. The central domain contains several signature residues of glycosyltransferases that are conserved in all species ( Richmond, 2000 ; Lei et al., 2012b ). Three conserved aspartic acid residues (D 1 , D 2 , and D 3 ) are distributed throughout the cytoplasmic domain and a QXXRW motif resides near the C-terminal end of the central domain ( Figure 2 ). These residues are often referred to collectively as the D,D,D,QXXRW motif and are thought to be involved in substrate binding, acceptor binding, and catalysis ( Saxena et al., 2001 ). In higher plants, CESAs have two extra plant-specific protein domains: a plant-conserved region (P-CR) that resides between D 1 and D 2 and a class-specific region (CSR) that resides between D 2 and D 3 The function of these two plant-specific domains is unclear, but it has been proposed that they might be involved in plant specific regulation of cellulose synthesis ( Sethaphong et al., 2013 ; Slabaugh et al., 2013 ).

Domain structure of cellulose synthase (CESA) protein and the positions of CESA mutations.

The direct proof that CESA is a component of the rosette TC came from an immunolabelling experiment in which freeze-fractured replicas from vascular plant Vigna angularis were labeled with CESA antibodies ( Kimura et al., 1999 ). Although the equivalent experiment has not been performed in Arabidopsis, a lesion in Arabidopsis CESA1 (AT4G32410) resulted in a deficiency in cellulose synthesis and caused a disintegration of rosettes, suggesting that CESA1 is a component of the rosette TC in Arabidopsis ( Arioli et al., 1998 ). The Arabidopsis genome encodes ten CESA genes ( Figure 2 and Table 1 ). Based on genetic analysis, CESAs are roughly classified into two groups that correspond to CESAs that are involved in cellulose synthesis in primary cell walls (primary CESAs ) and CESAs that are involved in cellulose synthesis in secondary cell walls (secondary CESAs ). Among primary CESAs , CESA1 and CESA3 (AT5G05170) are essential while CESA6 (AT5G64740) is redundant with CESA2 (AT4G39350), CESA5 (AT5G09870), and CESA9 (AT2G21770). CESA4 (AT5G44030), CESA7 (AT5G17420), and CESA8 (AT4G18780) are non-redundantly required for cellulose synthesis in secondary cell walls. The functional separation might reflect a need for distinct structural properties of CESA proteins to assemble different CSCs for primary and secondary cell wall synthesis. However, studies also indicate that the involvement of CESA genes in cellulose synthesis of primary and secondary cell walls could be flexible. For example, loss of function of CESA5 or CESA9 resulted in defects of seed coat formation, a process thought to be dependent upon cellulose synthesis in secondary walls ( Stork et al., 2010 ; Harpaz-Saad et al., 2011 ; Mendu et al., 2011b ; Mendu et al., 2011a ; Harpaz-Saad et al., 2012 ). Furthermore, ectopic overexpression of fra5 , a dominant mutant of CESA7 affects cell wall formation in both types of walls ( Zhong et al., 2003 ). In a genetic swap experiment, CESA7 , under the expression of the CESA3 promoter, can partially complement the growth defect of cesa3 je5 , indicating that CESA7 may have structural properties allowing its incorporation into primary CSCs ( Carroll et al., 2012 ). On the other hand, CESA1 , under a promoter of a secondary CESA gene, partially complemented cesa8 irx1 null mutant phenotypes, including defects in cellulose content, plant stature and collapsed xylem vessels ( Carroll et al., 2012 ; Li et al., 2013 ). Interestingly, a phylogenetic analysis revealed that the separation of primary and secondary CESAs may have occurred before the appearance of vascular plants ( Carroll et al., 2011 ).

Genes and mutations implicated in cellulose synthesis

3.2. Rosette assembly

Based on the genetic analysis of CESA proteins in Arabidopsis, an early model suggested that each of the six subunits of a rosette contains six CESA proteins, and that each rosette therefore has a total of 36 CESA proteins ( Scheible et al., 2001 ; Doblin et al., 2002 ). This model explains the formation of a rosette through the distinct binding properties among three different CESA proteins. However, this model has not been experimentally tested. The exact composition and stoichiometry of CESAs in a rosette is unclear. Without such information, the mechanism by which the rosette is assembled from individual CESA proteins remains a mystery. Transmission electron microscopy studies have visualized fully assembled rosettes in the Golgi apparatus ( Haigler et al., 1986 ). This is consistent with the observation that fluorescent protein-labeled CESAs accumulate in the Golgi apparatus, and suggests that rosettes are assembled prior to being delivered to the plasma membrane. Consistent with this idea, rosettes have also been observed in vesicles budding from the Golgi in Micrasterias denticulate ( Giddings et al., 1980 ). Because cellulose synthases are proteins with multiple transmembrane domains, it is reasonable to speculate that rosette assembly could occur at any stage in secretion as early as the endoplasmic reticulum or as late as the Golgi apparatus ( Wightman et al., 2010b ). In Arabidopsis, genetic studies of both primary and secondary cell walls indicated that at least three different CESAs are required for the formation of an intact CSC ( Doblin et al., 2002 ; Gardiner et al., 2003 ; Taylor et al., 2003 ). For secondary cell wall synthesis, deletion of CESA4, CESA7, or CESA8 resulted in a loss of rosette assembly and the remaining CESAs were not trafficked to cell wall deposition sites ( Taylor et al., 1999 ; Taylor et al., 2000 ; Gardiner et al., 2003 ; Taylor et al., 2003 ). Similarly, the temperature sensitive mutant, radial swelling root 1 (rsw1) caused the rosette to disintegrate, revealing the importance of CESA1 in the formation of an intact CSC during the synthesis of primary cell walls ( Arioli et al., 1998 ). Genetic data suggest that primary CSCs are composed of CESA1, CESA3, CESA6 or CESA6-like proteins. Since CESA1 and CESA3 are essential components in the CSC and CESA1 and CESA3 share the highest amino acid similarity, it is reasonable to speculate that CESA1 and/or CESA3 may reside in a special position in the CSC. A chimeric study of CESA1 and CESA3 indicated that these two CESAs have specific positions within the CSC ( Wang et al., 2006 ).

It has been speculated that the dimerization of CESA proteins is important for the formation of the rosette. Attempts to affinity purify an intact CSC using epitope-tagged CESAs led to the isolation of CESA oligomers ( Atanassov et al., 2009 ). The oligomers might be intermediates in the assembly of the complex. The zinc finger domain, the P-CR, and/or CSR domains are only present in rosette forming organisms ( Delmer, 1999 ), therefore, these domains are the obvious candidates for involvement in rosette assembly. An in vitro assay revealed that the zinc-binding domains of GhCESA1 and GhCESA2 were capable of forming homo-dimers and hetero-dimers ( Kurek et al., 2002 ). Interactions between CESAs have been demonstrated by co-immunoprecipitation experiments and yeast two-hybrid assays ( Taylor et al., 2003 ; Desprez et al., 2007 ; Timmers et al., 2009 ; Carroll et al., 2012 ; Li et al., 2013 ). A recent 3-D model of the central domain of CESA indicates that the P-CR and CSR domains may play a role in rosette assembly ( Sethaphong et al., 2013 ), however, it remains to be determined how the P-CR and/or CSR domains are involved in the interaction ( Carroll et al., 2012 ; Li et al., 2013 ).

3.3. Functions of the rosette

Formation of crystalline cellulose requires a simultaneous synthesis of multiple glucan chains through the plasma membrane rosettes. The rosettes are thought to have two distinct functions: intracellular polymerization of glucan chains and extracellular assembly of glucan chains into crystalline microfibrils ( Arioli et al., 1998 ). The exact number of glucan chains in an elementary cellulose microfibril remains controversial. The most popular model predicts that 36 glucan chains are synthesized simultaneously by one single rosette ( Scheible et al., 2001 ; Doblin et al., 2002 ). This prediction implies that 36 enzymatically active CESAs comprise each six-lobed rosette with each lobe being comprised of a hexamer of CESAs. It remains a challenge to express CESAs in vitro and test whether all of the ten CESAs in Arabidopsis are indeed enzymatically active. If some of the CESAs are not enzymatically active but rather acting as a scaffold, it poses an interesting situation in which the spacing of individual glucan chains may vary and may affect the crystallization process.

As of yet, a crystal structure of a CESA protein from higher plants has not been obtained. However, two recent studies have provided significant progress in revealing the structure of cellulose synthases. The first crystal structure of a cellulose synthase has been solved from the bacterium, Rhodobacter spaeroides , and the first predicted tertiary model of the central cytosolic domain of a plant CESA has been revealed ( Morgan et al., 2013 ; Sethaphong et al., 2013 ). The crystal structure of bacterial CESA, BcsA, supports the involvement of the D,D,D,QXXRW in catalysis and the idea that newly synthesized glucan chains are transported across the plasma membrane through a pore composed of CESA transmembrane domains. The bacterial crystal structure also provides important implications on CESA function in higher plants. In support of this claim, the predicted structure of the central domain of a cotton CESA is superimposable with the crystal structure of the bacterial CESA, BcsA. Consistent with these structural analyses, the Arabidopsis dominant negative mutant, thanatos (Pro578Ser), maps to the invariant QTPH sequence of bacterial BcsA, indicating that the catalytic base function of this amino acid might be conserved ( Daras et al., 2009 ; Morgan et al., 2013 ). The predicted tertiary structure of the cotton CESA suggests that the P-CR and the CSR domains of cotton CESA fold into distinct subdomains on the periphery of the catalytic region ( Morgan et al., 2013 ; Sethaphong et al., 2013 ). The putative functions of the P-CR and the CSR may include CSC subunit assembly and interactions with factors involved in the regulation of cellulose synthesis ( Sethaphong et al., 2013 ). In comparing the model of cotton CESA to Arabidopsis sequences, it was revealed that many of the existing Arabidopsis CESA mutation sites are located near the predicted catalytic region. For example, CESA1 E779K (rsw1-45) , CESA8 D683N (irx1-1) , CESA1 D780N (rsw1-20), and CESA7 D524N reside in the catalytic core ( Taylor et al., 2000 ; Beeckman et al., 2002 ; Liang et al., 2010 ; Sethaphong et al., 2013 )( Figure 2 ). While the validation of the model requires additional experiments, the model provides valuable insights into the mechanism of cellulose synthesis in higher plants.

Calcofluor disrupts the crystallization of cellulose in Acetobacter xylinum presumably by interfering with inter-chain hydrogen bonding. This disruption of cellulose microfibril assembly was accompanied by a four-fold increase in the rate of glucose polymerization ( Benziman et al., 1980 ; Haigler et al., 1980 ). These observations form the basis of a hypothesis that polymerization and crystallization are coupled processes during cellulose biosynthesis. It was proposed that crystallization occurs after the glucan chains have exited the cellulose synthase pores. The structure of BcsA-BcsB from Rhodobacter sphaeroides suggests that glucan synthesis and translocation is a coupled process in which the nascent glucan chain is extended by one glucose molecule at a time ( Morgan et al., 2013 ). If this structure holds true for Acetobacter xylinum , the rate of polymerization will be limited not only by the time required for glucan chains to bundle and crystallize but also by the rate of glucan translocation. Unlike bacteria in which the linear CSCs are stationary with respect to cell surface, rosette CSCs in plants and algae move within the plane of plasma membrane. In Arabidopsis, fluorescently tagged CESAs were observed to move through the plasma membrane at an average speed of 300–350 nm/min. The velocity of CSC movement through the plasma membrane provides a parameter by which the polymerization rate of glucan chains can be measured ( Paredez et al., 2006 ). Recently it was shown that CESA1 A903V and CESA3 T942I mutants displayed reduced crystalline cellulose content and crystallite size, accompanied by 16% and 8 % increase of velocity of CSCs, respectively ( Harris et al., 2012 ). Although the increase of reaction rate in plants might not be comparable to that of bacteria, these results provide evidence that plants may share a conserved mechanism to couple the polymerization rate with crystallization. Furthermore, overexpression of a cellulose-binding domain (CBD) in poplar resulted an increase in cellulose production, indicating that the crystallization process might also influence the polymerization rate in higher plants ( Levy et al., 2002 ).

Interestingly, the quinoxyphen-resistant mutant, CESA1 A903V , aligns with the Tyr455 residue in transmembrane domain 6 of BcsA, which forms a hydrogen bond to the translocating glucan, suggesting that quinoxyphen might affect the translocation of the glucan rather than the synthesis of the glucan ( Morgan et al., 2013 ). It is possible that this mutation affects the configuration of the nascent chain before crystallization, which leads to an affect on the crystallinity of the elementary microfibril and the size of the microfibril. Even if crystallization is a rate-limiting factor for glucan polymerization in higher plants, the difference in magnitude of the influence of crystallization on polymerization rate in bacteria compared to Arabidopsis suggests that regulatory mechanisms of the cellulose synthesis complex differ in the two evolutionarily distant species. Cellulose crystallization in bacteria occurs after glucan chains have been transported out of the cell ( Ross et al., 1991 ), whereas the crystallization of glucan chains in higher plants might be a function of the CSC. If polymerization and crystallization are indeed tightly coupled in cellulose biosynthesis in plants, the time required for glucan chains to bundle and crystallize may limit the rate of polymerization.

3.4. In vitro cellulose synthesis

Attempts to purify enzymatically active CSCs have been unsuccessful. As of yet, in vitro synthesis of cellulose using biochemical approaches has not been established in higher plants. The use of membrane fractions purified from plant tissues has demonstrated limited in vitro cellulose synthesis activities ( Okuda et al., 1993 ; Kudlicka et al., 1995 ; Kudlicka et al., 1997 ; Lai-Kee-Him et al., 2002 ; Ohlsson et al., 2006 ; Cifuentes et al., 2010 ). However, rather than cellulose, callose (β-1,3-glucan) appears to be the major product in these limited successful attempts. The disintegration of rosettes in rsw1 mutants indicates that production of crystalline cellulose may require an intact rosette ( Arioli et al., 1998 ). This may explain the difficulties in attempting to purify active complexes for in vitro cellulose synthesis as the purification of intact rosettes is challenging. It has been proposed that structural lipids of the plasma membrane may play an important role in keeping the CSCs intact ( Guerriero et al., 2010 ). The role of lipids in cellulose synthesis in vivo has been supported by the observation that lipids of the sterol family are crucial for cellulose synthesis and cell elongation ( Schrick, 2004 ). It is an attractive idea that the rosette is embedded in a membrane microdomain with a special lipid composition that is required for the integrity of the rosette and the regulation of cellulose synthesis ( Guerriero et al., 2010 ; Schrick et al., 2012 ). Nonetheless, a comprehensive characterization of all of the components necessary for maintaining the integrity and the activity of the rosette will be required to develop a successful in vitro cellulose synthesis assay in higher plants.

3.5. Genetic studies of CESA genes

Cellulose microfibrils are the major components in both primary and secondary cell walls. Therefore, the content and organization of cellulose microfibrils determine cell wall properties and ultimately control plant development. In Arabidopsis, cellulose deficiency in primary cell walls leads to a reduction in cell expansion ( Somerville, 2006 ; Lei et al., 2012b ) ( Figure 2 and Table 1 ). Mutations in either of the two essential CESA s, CESA1 or CESA3 , caused abnormal morphogenesis during embryo development ( Arioli et al., 1998 ; Beeckman et al., 2002 ; Gillmor et al., 2002 ; Daras et al., 2009 ). During post-embryonic development, the reduced cell expansion phenotype results in reduced cell elongation and exaggerated radial expansion in epidermal cells of dark grown hypocotyls and in roots of light grown seedlings ( Baskin, 2005 ; Crowell et al., 2010a ). For example, the first identified CESA mutation radial swelling root 1 (rswl) , a mutation of A589V in Arabidopsis CESA1 , showed an enhanced radial expansion defect at the restrictive temperature ( Arioli et al., 1998 ). A null mutant of CESA6 affected the normal cell morphogenesis in both root and etiolated hypocotyls ( Desnos et al., 1996 ; Fagard et al., 2000b ). Reverse genetic analyses revealed that the CESA6 like genes, including CESA2, CESA5, and CESA9 are collectively essential for cellulose synthesis ( Desprez et al., 2007 ; Persson et al., 2007 ). Interestingly, the loss of function mutant of CESA5 or CESA9 also altered seed coat mucilage structure ( Stork et al., 2010 ; Harpaz-Saad et al., 2011 ; Mendu et al., 2011b ; Mendu et al., 2011a ; Harpaz-Saad et al., 2012 ).

Defects in secondary wall cellulose synthesis are represented by collapsed or irregular xylem cells in the vasculature and reduced mechanical strength in fiber cells ( Turner et al., 1997 ). Irregular xylem (irx) mutants such as irx1-1, irx3 and irx5 encode CESA8, CESA7 and CESA4 , respectively ( Taylor et al., 1999 ; Taylor et al., 2000 ; Taylor et al., 2003 ). Irx1-1 has a mutation in aspartic acid 683 (D683N), which is highly conserved among all plant CESAs. Loss of function of any single secondary CESA causes a complete xylem morphology defect, indicating that each secondary CESA is important for proper function of the whole complex ( Taylor et al., 2000 ). Collapsed xylem vessels in several exigua (exi) mutants, which were mapped to CESA4, CESA7 , and CESA8 , caused a reduction in water transport capacity and reduced the cell expansion of cell types that do not form secondary cell walls, presumably due to turgor pressure defects ( Rubio-Diaz et al., 2012 ). Fragile fiber 5 (frab) , a dominant mutant of CESA7 , caused a reduction in cellulose content and thickness of the secondary wall of fiber cells ( Zhong et al., 2003 ). Cellulose synthesis inhibitors such as isoxaben and quinoxyphen have been used in genetic screens for resistant proteins ( Brabham et al., 2012 ). Two CESA mutants, ixr1 and ixr2 conferred resistance to isoxaben ( Scheible et al., 2001 ; Desprez et al., 2002 ) and a CESA3 mutant was resistant to quinoxyphen ( Harris et al., 2012 ).

Altered cellulose synthesis affects the physical and structural properties of the cell wall. Plant cells can sense the changes in the wall through cell wall integrity sensing mechanisms ( Wolf et al., 2012 ). For example, deficiency in cellulose synthesis in primary cesa mutants is often accompanied by excessive deposition of lignin in non-lignified cells ( Cano-Delgado et al., 2003 ; Hematy et al., 2007 ). Excessive lignification is also seen in etl1 (At1g05850) , a non -CESA mutant with defects in cellulose synthesis ( Zhong et al., 2002 ). Although the molecular mechanism by which a reduction in cellulose biosynthesis triggers the lignification is unknown, it has been speculated that it is part of a response to cell wall damage. Consistent with the cell wall damage hypothesis, lesions in primary CESAs induced accumulation of jasmonic acid and ethylene, two plant hormones that have a major role in plant defense, indicating that cellulose deficiency mimics the effect of physical damage or certain biotic stresses ( Ellis et al., 2001 ; Ellis et al., 2002 ; Cano-Delgado et al., 2003 ).

Inhibition of cellulose synthesis also alters plant hormone biosynthesis or signaling. A mutation in CESA3 (repp3) has been associated with altered PIN polarity that is essential for auxin flow in plants ( Feraru et al., 2011 ). Given the importance of auxin trafficking during embryogenesis, it is possible that the defects in embryogenesis seen in CESA1 mutants might, to some degree, result from altered auxin transport ( Beeckman et al., 2002 ). Mutations in all three secondary CESAs confer enhanced resistance to the bacterium, Ralstonia solanacearum , and the necrotrophic fungus, Plectosphaerella cucumerina , presumably through an abscisic acid (ABA) dependent pathway ( Chen et al., 2005 ; Hernandez-Blanco et al., 2007 ). Consistent with altered ABA synthesis, mutations in CESA8 (lew2) are more tolerant to drought stress and accumulate ABA ( Chen et al., 2005 ).

3.6. Non- CESA genes involved in cellulose synthesis

Through forward genetic screens, many non -CESA encoding genes have also been identified to be involved in cellulose synthesis in primary cell walls ( Hauser et al., 1995 ; Reiter et al., 1997 ; Nicol et al., 1998 ; Lane et al., 2001 ; Lukowitz et al., 2001 ; Burn et al., 2002 ; Gillmor et al., 2002 ; Pagant et al., 2002 ; Williamson et al., 2002 ; Scheible et al., 2003 ; Gillmor et al., 2005 ) ( Table 1 ). It remains to be determined whether any of the corresponding non-CESA proteins are directly or indirectly associated with CSCs. Nevertheless, these mutants indicate that cellulose synthesis in higher plants is an intricate and regulated process.

Some well-characterized cellulose deficient mutants include COBRA (AT5G60920), COBL4 (AT5G15630), POM-POM1 (AT1G05850), KOBITO1 (AT3G08550), and KORRIGAN1 (AT5G49720). COBRA (COB1) encodes a GPI-anchored extracellular protein ( Schindelman et al., 2001 ). cob1 disrupts cellulose microfibril orientation and COB1 localizes to microtubule-like structures. Therefore, COB1 has been proposed to be involved in the organization of cellulose microfibrils through a microtubule-related function ( Roudier et al., 2005 ). A COB1 paralog (COBL4/irx6) is involved in cellulose synthesis of secondary cell walls ( Brown et al., 2005 ). Mutations in POM-POM1/CTL1 , which encode a putative chitinase, cause dwarfism and cellulose deficiency ( Hauser et al., 1995 ; Zhong et al., 2002 ). CTL1 might affect cellulose synthesis through an interaction with cellulose ( Sanchez-Rodriguez et al., 2012 ). The KOBITO1/ELD1/ABI8 gene was isolated from screens for mutants with cellulose deficiency and dwarfism, and in screens for mutants with ABA insensitivity ( Pagant et al., 2002 ; Lertpiriyapong et al., 2003 ; Brocard-Gifford et al., 2004 ). Similar to the cob1 mutant, the cellulose microfibrils of kobito1 were mis-oreinted in epidermal cells of root elongation zone ( Pagant et al., 2002 ).

Lesions in KORRIGAN1 (KOR1 ) resulted in defects in cellulose synthesis in both primary and secondary cell walls, including defects in cytokinesis and unrestricted cell proliferation, root radial swelling, dwarfism and collapsed xylem ( Nicol et al., 1998 ; Zuo et al., 2000 ; Lane et al., 2001 ; Sato et al., 2001 ; Szyjanowicz et al., 2004 ; Paredez etal., 2008 ). KOR1 encodes a putative membrane bound β-1,4 endoglucanase ( Nicol et al., 1998 ; Rudsander et al., 2008 ; Liebminger et al., 2013 ). KOR1 from Brassica napus that was heterologously expressed and purified in Pichia pastoris was able to digest non-crystalline cellulose but not crystalline cellulose or xyloglucan ( Molhoj et al., 2001 ). Similar enzymatic activity has been observed in a KOR1 ortholog from poplar ( Master et al., 2004 ). GFP-KOR1 was localized to the Golgi apparatus, endosomal compartments and the plasma membrane in Arabidopsis ( Robert et al., 2005 ; Crowell et al., 2010b ). In bacteria and Rhizobium, a cellulase gene is part of the cellulose synthase operon and plays an essential role in cellulose synthesis ( Hayashi et al., 2005 ). KOR1 has several proposed functions including: cleaving of a sitosterol-glucoside, a putative primer for cellulose synthesis initiation ( Molhoj et al., 2002 ; Peng et al., 2002 ); removing or modifying erroneous glucan chains; and releasing the CSCs from the growing glucan chains at the end of cellulose microfibrils ( Nicol et al., 1998 ; Delmer, 1999 ). These proposed functions might suggest a direct association exists between KOR1 and CSCs. However, localization studies indicate that KOR1 and CSCs lack a tight association ( Szyjanowicz et al., 2004 ). Immunoprecipitation experiments have also failed to prove a direct interaction between KOR1 and CESAs ( Desprez et al., 2007 ). When the KOR1 ortholog, PttCel9A, was superimposed onto the crystal structure of a homologous bacterial enzyme, it revealed key differences in the active sites ( Master et al., 2004 ). The lack of a few key determinants for substrate binding may explain why the poplar ortholog has such low catalytic activity. The Arabidopsis root-swelling mutant, rsw2, which is a KOR1 allele, has a defect in crystalline cellulose production at restrictive temperatures, a phenotype that is similar to that of rswl ( Lane et al., 2001 ). Together with the Golgi localization of GFP-KOR1, it might be possible that KOR1 is important for maintaining an intact rosette. Overall, KOR1 might be a multi-functional protein in regulating cellulose synthesis.

Similar to cesal and cesa3 mutants, lesions in enzymes involved in N-glycosylation show embryo defects and cellulose deficiency ( Lukowitz et al., 2001 ; Gillmor et al., 2002 ). Although CESA proteins have not been reported to be N-glycosylated, it is possible that N-glycan modification is required for proteins that are functionally associated with CSCs. For example, Arabidopsis KOR1 is subjected to N-glycosylation at several positions and evidence from in vitro studies with KOR1 and its orthologs from Brassica napus have shown that N-glycan modification of KOR1 is important for its cellulase activity ( Molhoj et al., 2001 ; Liebminger et al., 2013 ).

Mutants with changes in sterol content also have cellulose content deficiencies ( Schrick, 2004 ). It has been proposed that sterol-glucoside acts as a primer that is required for cellulose synthesis ( Peng et al., 2002 ). In addition to being a primer, sterols may also be involved in the targeting of CSCs to the plasma membrane and in maintaining rosette integrity and activity ( Schrick et al., 2012 ).

4. REGULATION OF CELLULOSE SYNTHESIS

4.1. regulation at the transcriptional level.

CESAs that are essential for primary cell wall synthesis are considered housekeeping genes since every plant cell is surrounded by a primary cell wall. Consistent with the this idea, CESA1, CESA3 and CESA6 transcripts are accumulated at high levels ubiquitously ( Hamann et al., 2004 ). The co-expression of primary CESAs and secondary CESAs are consistent with the hypothesis that CSCs consist of at least three isoforms for cellulose synthesis in primary and secondary cell wall ( Brown et al., 2005 ; Persson et al., 2005 ). A transgenic line in which the CESA6 promoter drives the expression of CESA2 cDNA complements a cesa6 prc mutant phenotype, suggesting that CESA2 is capable of replacing CESA6 in the CSC. Further supporting the transcriptional coregulation of CESAs, the transcription of CESA2 and CESA6 was regulated by ethylene in a similar fashion ( Hamann etal., 2004 ).

Hormones can have various effects on the expression of CESAs . For example, brassinosteroid (BRs) biosynthesis or perception deficiency is accompanied with defects in cellulose synthesis ( Schrick et al., 2012 ). Exogenous application of BR recovers the deficiency of CESA expression in the BR biosynthesis mutatnt, det2-1 (AT2G38050), but not in the BR receptor mutant, bri1-301 (AT4G39400), suggesting that BR signaling is important for transcriptional regulation of cellulose biosynthesis ( Xie et al., 2011 ). Consistent with this hypothesis, BES1 (AT1G19350), a BR-activated transcription factor was shown to interact with upstream elements of many CESA genes ( Xie et al., 2011 ).

Transcriptional and genetic analyses have identified a cascade of transcriptional events that are critical for secondary wall synthesis initiation ( Zhong et al., 2010 ). NAC domain transcription factors such as SECONDARY WALL-ASSOCIATED NAC DOMAIN PROTEIN (SND1 ) (AT1G32770) and NAC SECONDARY WALL THICKENING PROMOTING FACTOR (NST1) (AT2G46770) are key regulators of the transcriptional cascade ( Zhong et al., 2006 ; Mitsuda et al., 2007 ). Double mutants of SND1 and NST1 resulted in a loss of secondary wall formation in the fibers of stems, which included a lack of cellulose ( Zhong et al., 2007a ). MYB46 (AT5G12870) and MYB83 (AT3G08500) transcription factors are direct targets of NAC regulators ( Zhong et al., 2007b ; Zhong et al., 2008 ; McCarthy et al., 2009 ). Over-expression of MYB46 or MYB83 increased the expression of secondary CESA genes and induced secondary cell wall deposition ( Ko et al., 2009 ; McCarthy et al., 2009 ). MYB46 can directly bind to promoters of the three secondary CESA genes as demonstrated by in vitro and in vivo assays ( Kim etal., 2012b ). Two studies identified consensus sequences for MYB binding, named the secondary wall MYB-responsive element (SMRE) or the MYB46-responsive cis-regulatory element (M46RE), which have been shown to be present in the promoter regions of many secondary cell wall related genes ( Zhong et al., 2012 ) ( Kim et al., 2012a ). The presence of multiple copies of M46RE in the three secondary CESA genes suggests that there is a direct transcriptional regulation of CESAs through MYB46. Genetic complementation experiments confirmed that M46REs are critical for cellulose synthesis in secondary cell walls ( Kim et al., 2013 ).

4.2. Regulation at the post-transcriptional level

The regulation of CESA at the post-transcriptional level may include the regulation of CSC abundance, the lifetime of the CSC at the plasma membrane, and the post-transcriptional modification of the CSC. The abundance and lifetime of CSCs in the plasma membrane may affect the amount and structural properties of cellulose microfibrils. Studies suggest that CESA proteins may have a short lifetime in the plasma membrane. For example, it was estimated that the lifetime of cotton CESA was less than 30 minutes ( Jacob-Wilk et al., 2006 ). CESA dimerization via the N-terminal zinc-binding domain is regulated by redox state and may affect the degradability of CESAs. Cotton GhCESA1 forms a dimer under oxidative conditions and is more resistant to degradation than the reduced monomeric form of GhCESA1, suggesting the N-terminal cytosolic portion of higher plant CESA proteins may be important for the stability of CESAs ( Kurek et al., 2002 ). Even though the zinc-binding domain is conserved among all CESAs from higher plants, it remains to be tested whether other CESAs, especially primary CESAs, are regulated in this manner.

Post-transcriptional modification may affect the stability and/or activity of CESAs. The phosphorylation of CESAs has attracted much attention because of the recent development in phosphoproteomic studies and genetic manipulation of CESAs. Phosphoproteomic studies in Arabidopsis and maize discovered that CESA proteins including CESA1, CESA3, and CESA5 are phosphorylated in conserved residues of the hypervariable region at the N-terminus ( Nuhse et al., 2004 ; Facette et al., 2013 ). The role of phosphorylation of CESA1 was tested by complementing the rsw1 mutant with mutated forms of CESA1 that prevent or mimic phosphorylation. Six phosphorylation sites were mutated either to alanine (A) to prevent phosphorylation or to glutamic acid (E) to mimic phosphorylation. The complementation was assessed by rescue of cell elongation, of cellulose content, and of CSC dynamics ( Chen et al., 2010 ). This study suggested that phosphorylation at T166, S686, and S688 is important for normal cellulose content and proper CSC velocity. Interestingly, a few of the single amino acid CESA1 phosphorylation-mutants caused asymmetric motility of CSCs in rsw1 mutant cells. The asymmetric movement appears to rely on an intact cortical microtubule structure since treatment with oryzalin, a microtubule depolymerizing drug, abolished the asymmetric movement ( Chen et al., 2010 ). The role of CESA5 phosphorylation has also been investigated. CESA5 is presumably redundant with CESA6. While the native form of CESA5 did not rescue cesa6 prc1-1 , a phosphorylation-mimicking CESA5 mutant with four serine residue substitutions was able to partially complement cesa6 prc1-1 ( Bischoff et al., 2011 ). Phosphorylation of CESA7 has been documented in vivo at two serine residues within a non-conserved region. These two serine residues were phosphorylated by plant extracts, resulting in protein degradation via a proteasome dependent pathway ( Taylor, 2007 ). While the molecular mechanism of CESA phosphorylation remains to be characterized, it is clear that phosphorylation of CESAs plays multiple roles in regulating the stability and activity of the CSC as well as the relationship between the CSC and microtubules.

4.3. Localization and trafficking of the CSC

CSCs were observed in various locations including the plasma membrane, Golgi-derived vesicles, and Golgi cisternae via transmission electron microscopy of freeze-fracture replicas ( Haigler et al., 1986 ). Observing the dynamic behavior of CSCs in living cells was not possible until the recent development of live imaging and fluorescent protein tagging of individual CESAs. Fusions between GFP variants and several CESA subunits, namely CESA3, 5, 6, and 7, did not interfere with the protein function as demonstrated by complementation of the respective mutant phenotypes with the fluorescent fusions ( Gardiner et al., 2003 ; Paredez et al., 2006 ; Desprez et al., 2007 ; Bischoff et al., 2011 ). Presumably, multiple copies of each fluorescent CESA isoform reside within a single CSC, allowing for the detection of individual or multiple CSCs within each diffraction-limited particle at the plasma membrane. Consistent with earlier freeze fracture studies in which rosettes were detected within the periphery of the trans face of the Golgi, fluorescent protein tagged CESAs also localize to the periphery of the Golgi apparatus ( Haigler et al., 1986 ; Crowell et al., 2009 ; Gutierrez et al., 2009 ). Immunogold labeling of a GFP-CESA3 fusion showed GFP-CESA3 localized to the ends of medial and trans-Golgi cisternae ( Crowell et al., 2009 ). The presence of CSC containing vesicles moving beneath the sites of secondary wall deposition has been inferred from fluorescence loss in photobleaching experiments ( Wightman et al., 2009 ). For primary cell wall synthesis, CSC-containing vesicles may reside in small subcellular compartments named small CESA compartments (SmaCCs) or microtubule associated cellulose synthase compartments (MASCs) ( Crowell et al., 2009 ; Gutierrez et al., 2009 ). Observations of SmaCCs/MASCs during cell expansion and under certain stress treatments indicate that they may function as storage vesicles ( Gutierrez et al., 2009 ). SmaCCs/MASCs accumulate in epidermal cells at the base of hypocotyl where cell expansion has ceased and CSC density is low ( Crowell et al., 2009 ). Osmotic stress and certain drug treatments deplete CSCs from the plasma membrane, which may help to reveal SmaCCs/ MASCs and/or induce SmaCC/MASC formation. The function of SmaCCs/MASCs has been interpreted in two different ways. Either SmaCCs/MASCs function as delivery compartments that appear before the insertion of CSCs into the plasma membrane or SmaCCs/MASCs function as intracellular storage vesicles of internalized CSCs. Upon drug treatment, rapid depletion of CSCs from the plasma membrane corresponds with an apparent accumulation of SmaCCs/MASCs, suggesting that SmaCCs/MASCs might result from the internalization of CSCs ( Crowell et al., 2009 ; Gutierrez et al., 2009 ; Wightman et al., 2010a ). SmaCCs/MASCs have been observed to be associated to cortical microtubules and track depolymerizing microtubule ends, which might be consistent with SmaCC/MASC involvement in assisting the delivery of CSCs along the cortical microtubules ( Paredez et al., 2006 ; Gu et al., 2010b ; Gu et al., 2010a ; Lei et al., 2012a ; Li et al., 2012c ). Alternatively, the association of SmaCCs/MASCs with microtubules might be related to the active transport of vesicles via microtubule motors ( Gutierrez et al., 2009 ).

In primary cell wall synthesizing epidermal cells, disruption of actin using Latrunculin B, an actin-destabilizing drug, results in reduced velocity of Golgi movement and aggregation of CSC containing Golgi bodies ( Gutierrez et al., 2009 ). As a result, a much higher density of plasma membrane-localized CSCs accumulates in regions directly above the aggregated Golgi bodies than in regions lacking underlying Golgi bodies ( Gutierrez et al., 2009 ). This suggests that actin is not crucial for the delivery of CSCs from the Golgi to the plasma membrane but that actin is important for the global distribution of CSCs. Although cortical microtubules appear to mark the sites of CSC delivery in primary cell walls, the depletion of microtubules did not affect the delivery rate of CSCs, suggesting that the delivery of CSCs is independent of microtubule function ( Gutierrez et al., 2009 ). In developing xylem cells, patterning of secondary cell wall deposition is characterized by bands of fluorescently labeled CSCs beneath the sites of cellulose deposition ( Wightman et al., 2008 ). In this system, YFP-CESA7 labeled Golgi bodies exhibit rapid movement along thick actin cables and a loss of actin bundles results in the cessation of Golgi movement and the loss of CSCs bands, suggesting that the actin cytoskeleton is required for the delivery of CSCs during secondary cell wall formation ( Wightman et al., 2008 ). It appears that the removal of microtubules does not affect the pausing of Golgi bodies, an event that is presumably associated with the delivery of CSCs at secondary cell wall thickening sites ( Wightman et al., 2008 ). It remains to be addressed whether Goklgi-mediated CSC insertion is responsible for the delivery of de novo CSCs, recycled CSCs or both ( Wightman et al., 2010b ; Lei et al., 2012b ).

Clathrin-mediated endocytosis (CME) is a well-characterized process by which eukaryotes internalize material from the plasma membrane and has recently been implicated in the internalization of CESAs from the plasma membrane ( Bashline et al., 2013 ). In CME, the adaptor protein complex 2 (AP2) acts in recruiting plasma membrane protein cargo and clathrin triskelia of the clathrin coat to the sites of endocytic vesicle formation ( Chen et al., 2011 ). µ2 (At5g46630), also referred to as AP2M, is a subunit of the AP2 complex in Arabidopsis ( L. Segal, 1959 ; Li et al., 2012a ; Bashline et al., 2013 ; Yamaoka et al., 2013 ). Live cell imaging analysis confirmed that the dynamics of µ2 is associated with CME in Arabidopsis ( Bashline et al., 2013 ). This study also demonstrated that CESA is a cargo protein of CME by displaying a direct interaction between CESAs and µ2 in vitro , by observing the participation of µ2-YFP in the endocytosis of CESA6-mCherry in planta , and by revealing that the abundance of YFP-CESA6 labeled CSCs at the plasma membrane is dependent on µ2. This study suggests that only a small fraction of CSCs are undergoing internalization at any given time ( Bashline et al., 2013 ).

The idea that CSCs are internalized from the plasma membrane through CME was challenged by the sizes of cytoplasmic portion of CSCs compared to the size of CME-derived vesicles ( Bowling et al., 2008 ; Crowell et al., 2009 ; Wightman et al., 2010a ; Bashline et al., 2011 ). The cytoplasmic portion of the CSC was estimated to be 45–50 nm in width ( Bowling et al., 2008 ) while the diameter of CME-derived vesicles without the clathrin coat was estimated to be an average of 30 nm in Arabidopsis ( Dhonukshe et al., 2007 ; Li et al., 2012a ). However, it is quite possible that CSCs may become structurally defective before being internalized and therefore CSCs that are undergoing internalization may contain a reduced number of subunits and have a reduced overall size. Alternatively, the measurements of the cytoplasmic portion of the CSC via TEM may be skewed by the association of accessory or regulatory proteins that bind to the CSC when it is functioning, but that dissociate from the CSC prior to the internalization of the CSC ( Bowling et al., 2008 ).

4.4. Microtubule cytoskeleton and CSC

The microtubule cytoskeleton is essential for controlling cell morphogenesis in higher plants ( Fisher et al., 1998 ; Burk et al., 2001 ; Burk et al., 2002 ; Paradez et al., 2006 ; Hamant et al., 2008 ; Boudaoud et al., 2009 ). The anisotropic growth of most post mitotic plant cells is determined by the transverse orientation (perpendicular to the axis of elongation) of cellulose microfibrils. An intimate relationship between microtubules and cellulose microfibrils was postulated at the time of the discovery of the microtubule cytoskeleton. In 1962, Green hypothesized that cytoplasmic elements, later named microtubules, at the cell periphery determined the orientation of newly synthesized cellulose microfibrils, and thereby determined the final shape of plant cells ( Green, 1962 ; Ledbetter et al., 1963 ). The parallel alignment between the orientation of cellulose microfibrils and the underlying cortical microtubules forms the basis of the “alignment hypothesis” ( Ledbetter et al., 1963 ). Many cytological data support the alignment hypothesis, although the microtubules do not predict the orientation of cellulose microfibrils in some cases ( Hepler et al., 1964 ; Baskin, 2001 ; Himmelspach et al., 2003 ; Sugimoto et al., 2003 ; Wasteneys, 2004 ; Wasteneys et al., 2004 ). Two molecular mechanisms have been put forward to explain the alignment hypothesis. In the “direct hypothesis”, CSCs are thought to be attached to cortical microtubules via a direct or indirect interaction, which causes the movement of CSCs and the synthesis of cellulose to be guided by microtubules ( Heath, 1974 ). In an alternative model, the “bumper hypothesis,” cellulose synthesis is constrained spatially by the presence of cortical microtubules that define channels within which CSCs can move at the plasma membrane without a physical link between the CSC and the microtubules ( Giddings et al., 1991 ). Recent live imaging provides convincing evidence to support the direct hypothesis. By dual labeling microtubules and CSCs using different fluorescent proteins, CSCs were observed to move along the underlying track provided by the cortical microtubules ( Paredez et al., 2006 ) ( Figure 3 ). The association between CSCs and the cortical microtubules was further demonstrated by the concomitant reorientation of both CSC trajectories and cortical microtubules upon blue light excitation and oryzalin treatment ( Paredez et al., 2006 ). While the live imaging of the intimate association between CSCs and microtubules rules out the bumper model, it does not differentiate between CSCs interacting with microtubules directly or through an indirect mechanism via other microtubule associated proteins (MAPs).

Cellulose synthase complexes (CSCs) and microtubules. Confocal images of epidermal cells of dark grown Arabidopsis seedlings expressing both YFP-CESA6 and RFP-TUA5. The coalignment of CSC trajactories and microtubules is evident in merged image. Time averaged image (duration 5 min, 5 sec interval) Scale bar = 10 mm.

To identify putative MAPs that facilitate the association between CSCs and microtubules, a yeast two-hybrid screen was performed using the central domain of primary CESAs ( Gu et al., 2010b ; Gu et al., 2010a ). Cellulose synthase interacting protein 1 (CSI1) (AT2G22125) was identified as an interaction partner of the central domain of CESA6. CSI1 was shown to fit the criteria for a CSC-microtubule linker protein in that CSI1 directly interacted with CESAs in a yeast two-hybrid assay and CSI1 directly interacted with microtubules in vitro ( Baskin et al., 2012 ). In addition, fluorescent protein-tagged CSI1 co-localized with CSCs in planta and exhibited bidirectional motility that was identical to that of CSCs, indicating that CSI1 is associated with CSCs in vivo ( Bringmann et al., 2012 ; Li et al., 2012c ). Also supporting the role of CSI1 as a linker protein, csi1 null mutants disrupted the association between CSCs and cortical microtubules in vivo ( Lei et al., 2012a , b ; Li et al., 2012c ).

In addition to mediating the interaction between CSCs and microtubules, CSI1 also is required to maintain the normal velocity of CSCs. Microtubules have been shown to modulate the velocity of CSCs. The velocity of fluorescently labeled CESA5 in cesa6 prc1-1 background increases after the depletion of cortical microtubules upon oryzalin treatment ( Bischoff et al., 2011 ). Prolonged treatment with oryzalin reduced the velocity of GFP-CESA6 labeled CSCs ( Li et al., 2012c ). It has also been reported a shorter oryzalin treatment at a low concentration did not significantly affect the velocity of CSCs containing GFP-CESA3 or YFP-CESA6, indicating that cortical microtubule structure is not solely responsible for maintaining normal activity of CSCs ( DeBolt et al., 2007 ; Chan et al., 2010 ). Loss of CSI1 resulted in reduced velocity of CSCs, which is comparable to the loss of microtubules upon prolonged treatment of oryzalin ( Brown, 1999b ; Gu et al., 2010b ; Gu et al., 2010a ). It remains to be determined whether CSI1-mediated activity of CSCs is a microtubule dependent process.

The alignment hypothesis might be a simplified model of a more complex association between cortical microtubules and cellulose synthesis. It is more likely that microtubules have additional functions in regulating cellulose biosynthesis. For example, microtubules may regulate exocytosis via direct transport of vesicles containing CSCs and their associated proteins such as KOR1 ( Robert et al., 2005 ; Fujita et al., 2012 ). Cortical microtubules have been proposed to change the composition of lipids in the plasma membrane, which may influence cellulose synthesis ( Fujita et al., 2012 ). It has also been observed that cortical microtubules compartmentalize endocytosis sites, a process capable of changing the physiological status of the plasma membrane ( Kakar et al., 2013 ). Overall, cortical microtubules most likely act as a platform for regulating many aspects of CSC related functions.

5. CONCLUDING REMARKS AND FUTURE DIRECTIONS

An understanding of the mechanism of cellulose synthesis in higher plants is important not only due to the vast everyday use of products that are made from cellulosic plant material, but also due to the importance of cellulose biosynthesis in plant development. Decoding the mechanism of cellulose synthesis in higher plants should be a pivotal part of understanding plant evolution. Cellulosic biofuel, an environmentally sustainable energy source, has the potential to significantly change the global energy supply and to reduce our dependency on fossil fuels. The future for economically adopting such biofuel is dependent on how well the cellulose biosynthesis process is understood ( Somerville, 2007 ; Carroll et al., 2009 ; Pauly et al., 2010 ). Arabidopsis has provided its wealth of genetic capability for understanding this process, however many fundamental questions remain to be addressed. For example, how is the rosette CSC assembled? Is there a primer for cellulose synthesis? How is CSC activated in the plasma membrane? What determines the lifetime of a CSC in the plasma membrane? What is the force/energy to power the movement of CSCs and how does it relate to mutants with defects in cellulose synthesis? How is crystallization controlled considering that the CSC is oriented perpendicular to the orientation of nascent microfibrils? How do cellulose elementary microfibrils form bundles? Many of these questions need innovations in the areas of cell biology, biochemistry, biophysics and computational modeling.

ACKNOWLEDGEMENTS

This work is supported by grant from National Science Foundation (1121375).

Citation: Shundai Li, Logan Bashline, Lei Lei and Ying Gu. (2014) Cellulose Synthesis and Its Regulation. The Arabidopsis Book 11:e0169. doi:10.1199/tab.0169

elocation-id: e0169

First published on January 13, 2014: e0169. doi: 10.1199/tab.0169

• Anderson C.T., Carroll A., Akhmetova L., Somerville C. Real-time imaging of cellulose reorientation during cell wall expansion in Arabidopsis roots. Plant Physiol. 2010; 152 :787–796. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Arioli T., Peng L., Betzner A.S., Burn J., Wittke W., Herth W., Camilleri C., Hofte H., Plazinski J., Birch R., Cork A., Glover J., Redmond J., Williamson R.E. Molecular analysis of cellulose biosynthesis in Arabidopsis. Science. 1998; 279 :717–720. [ PubMed ] [ Google Scholar ]
• Atalla R.H., Vanderhart D.L. Native Cellulose - a Composite of 2 Distinct Crystalline Forms. Science. 1984; 223 :283–285. [ PubMed ] [ Google Scholar ]
• Atanassov II, Pittman. J.K. Turner S.R. Elucidating the mechanisms of assembly and subunit interaction of the cellulose synthase complex of Arabidopsis secondary cell walls. J. Biol. Chem. 2009; 284 :3833–3841. [ PubMed ] [ Google Scholar ]
• Barnett J.R., Bonham V.A. Cellulose microfibril angle in the cell wall of wood fibres. Biol. Rev. Cam. Philo. Soc. 2004; 79 :461–472. [ PubMed ] [ Google Scholar ]
• Barnette A.L., Lee C., Bradley L.C., Schreiner E.P., Park Y.B., Shin H., Cosgrove D.J., Park S., Kim S.H. Quantification of crystalline cellulose in lignocellulosic biomass using sum frequency generation (SFG) vibration spectroscopy and comparison with other analytical methods. Carbo. Polymers. 2012; 89 :802–809. [ PubMed ] [ Google Scholar ]
• Bashline L., DU J., Gu Y. The trafficking and behavior of cellulose synthase and a glimpse of potential cellulose synthesis regulators. Front. Biol. 2011; 6 :377–383. [ Google Scholar ]
• Bashline L., Li S., Anderson C.T., Lei L., Gu Y. The endocytosis of cellulose synthase in Arabidopsis is dependent on mu2, a clathrin mediated endocytosis adaptin. Plant Physiol; 2013. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Baskin T.I. On the alignment of cellulose microfibrils by cortical microtubules: a review and a model. Protoplasma. 2001; 215 :150–171. [ PubMed ] [ Google Scholar ]
• Baskin T.I. Anisotropic expansion of the plant cell wall. Annu. Rev. Cell Dev. Biol. 2005; 21 :203–222. [ PubMed ] [ Google Scholar ]
• Baskin T.I., Gu Y. Making parallel lines meet: transferring information from microtubules to extracellular matrix. Cell Adhesion Migration. 2012; 6 :404–408. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Beeckman T., Przemeck G.K., Stamatiou G., Lau R., Terryn N., De Rycke R., Inze D., Berleth T. Genetic complexity of cellulose synthase a gene function in Arabidopsis embryogenesis. Plant Physiol. 2002; 130 :1883–1893. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Benziman M., Haigler C.H., Brown R.M., White A.R., Cooper K.M. Cellulose biogenesis: Polymerization and crystallization are coupled processes in Acetobacter xylinum. Proc. Natl. Acad. Sci. USA. 1980; 77 :6678–6682. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Bichet A., Desnos T., Turner S., Grandjean O., Hofte H. BOTERO1 is required for normal orientation of cortical microtubules and anisotropic cell expansion in Arabidopsis. Plant J. 2001; 25 :137–148. [ PubMed ] [ Google Scholar ]
• Bischoff V., Desprez T., Mouille G., Vernhettes S., Gonneau M., Hofte H. Phytochrome Regulation of Cellulose Synthesis in Arabidopsis. Curr. Biol. 2011; 21 :1822–1827. [ PubMed ] [ Google Scholar ]
• Bootten T.J., Harris P.J., Melton L.D., Newman R.H. Using Solid-State C-13 NMR Spectroscopy to Study the Molecular Organisation of Primary Plant Cell Walls. Plant Cell Wall: Methods and Protocols. 2011; 715 :179–196. [ PubMed ] [ Google Scholar ]
• Bosca S., Barton C.J., Taylor N.G., Ryden P., Neumetzler L., Pauly M., Roberts K., Seifert G.J. Interactions between MUR10/CesA7-dependent secondary cellulose biosynthesis and primary cell wall structure. Plant Physiol. 2006; 142 :1353–1363. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Boudaoud A., Corson F., Hamant O., Bohn S., Traas J., Couder Y. Turning a plant tissue into a living cell froth through isotropic growth. Proc. Natl. Acad. Sci. USA. 2009; 106 :8453–8458. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Bowling A.J., Brown R.M. The cytoplasmic domain of the cellulose-synthesizing complex in vascular plants. Protoplasma. 2008; 233 :115–127. [ PubMed ] [ Google Scholar ]
• Brabham C., Debolt S. Chemical genetics to examine cellulose biosynthesis. Front. Plant Sci. 2012; 3 :309. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Brett C.T. Cellulose microfibrils in plants: biosynthesis, deposition, and integration into the cell wall. Intl. Rev. Cytol. 2000; 199 :161–199. [ PubMed ] [ Google Scholar ]
• Bringmann M., Li E., Sampathkumar A., Kocabek T., Hauser M.T., Persson S. POM-POM2/cellulose synthase interacting1 is essential for the functional association of cellulose synthase and microtubules in Arabidopsis. Plant Cell. 2012; 24 :163–177. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Brocard-Gifford I., Lynch T.J., Garcia M.E., Malhotra B., Finkelstein R.R. The Arabidopsis thaliana ABSCISIC ACID-INSENSITIVE8 encodes a novel protein mediating abscisic acid and sugar responses essential for growth. Plant Cell. 2004; 16 :406–421. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Brown D.M., Zeef L.A., Ellis J., Goodacre R., Turner S.R. Identification of novel genes in Arabidopsis involved in secondary cell wall formation using expression profiling and reverse genetics. Plant Cell. 2005; 17 :2281–2295. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Brown R.M. Cellulose structure and biosynthesis. Pure and Applied Chemistry. 1999a; 71 :767–775. [ Google Scholar ]
• Brown R.M. Cellulose structure and biosynthesis: Where do we stand today?. Abstracts of Papers of the American Chemical Society. 1999b; 218 :U214–U214. [ Google Scholar ]
• Brown R.M., Saxena I.M. Cellulose biosynthesis: A model for understanding the assembly of biopolymers. Plant Physiol. Biochem. 2000; 38 :57–67. [ Google Scholar ]
• Brown R.M., Saxena I.M., Kudlicka K. Cellulose biosynthesis in higher plants. Trends Plant Sci. 1996; 1 :149–156. [ Google Scholar ]
• Burk D.H., Ye Z.H. Alteration of oriented deposition of cellulose microfibrils by mutation of a katanin-like microtubule-severing protein. Plant Cell. 2002; 14 :2145–2160. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Burk D.H., Liu B., Zhong R., Morrison W.H., Ye Z.H. A katanin-like protein regulates normal cell wall biosynthesis and cell elongation. Plant Cell. 2001; 13 :807–827. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Burn J.E., Hurley U.A., Birch R.J., Arioli T., Cork A., Williamson R.E. The cellulose-deficient Arabidopsis mutant rsw3 is defective in a gene encoding a putative glucosidase II, an enzyme processing N-glycans during ER quality control. Plant J. 2002; 32 :949–960. [ PubMed ] [ Google Scholar ]
• Cano-Delgado A., Penfield S., Smith C., Catley M., Bevan M. Reduced cellulose synthesis invokes lignification and defense responses in Arabidopsis thaliana. Plant J. 2003; 34 :351–362. [ PubMed ] [ Google Scholar ]
• Carroll A., Somerville C. Cellulosic Biofuels. Annu. Rev. Plant Biol. 2009; 60 :165–182. [ PubMed ] [ Google Scholar ]
• Carroll A., Specht C.D. Understanding Plant Cellulose Synthases through a Comprehensive Investigation of the Cellulose Synthase Family Sequences. Front. Plant Sci. 2011; 2 :5. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Carroll A., Mansoori N., Li S.D., Lei L., Vernhettes S., Visser R.G.F., Somerville C., Gu Y., Trindade L.M. Complexes with Mixed Primary and Secondary Cellulose Synthases Are Functional in Arabidopsis Plants. Plant Physiol. 2012; 160 :726–737. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Chaffey N. Wood formation in forest trees: from Arabidopsis to Zinnia. Trends Plant Sci. 1999; 4 :203–204. [ PubMed ] [ Google Scholar ]
• Chan J., Calder G., Fox S., Lloyd C. Cortical microtubule arrays undergo rotary movements in Arabidopsis hypocotyl epidermal cells. Nat. Cell Biol. 2007; 9 :171–175. [ PubMed ] [ Google Scholar ]
• Chan J., Crowell E., Eder M., Calder G., Bunnewell S., Findlay K., Vernhettes S., Hofte H., Lloyd C. The rotation of cellulose synthase trajectories is microtubule dependent and influences the texture of epidermal cell walls in Arabidopsis hypocotyls. J. Cell Sci. 2010; 123 :3490–3495. [ PubMed ] [ Google Scholar ]
• Chen S., Ehrhardt D.W., Somerville C.R. Mutations of cellulose synthase (CESA1) phosphorylation sites modulate anisotropic cell expansion and bidirectional mobility of cellulose synthase. Proc Natl Acad Sci U S A. 2010; 107 :17188–17193. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Chen X., Irani N.G., Friml J. Clathrin-mediated endocytosis: the gateway into plant cells. Curr. Opin. Plant Biol. 2011; 14 :674–682. [ PubMed ] [ Google Scholar ]
• Chen Z., Hong X., Zhang H., Wang Y., Li X., Zhu J.K., Gong Z. Disruption of the cellulose synthase gene, AtCesA8/IRX1, enhances drought and osmotic stress tolerance in Arabidopsis. Plant J. 2005; 43 :273–283. [ PubMed ] [ Google Scholar ]
• Cifuentes C., Bulone V., Emons A.M.C. Biosynthesis of Callose and Cellulose by Detergent Extracts of Tobacco Cell Membranes and Quantification of the Polymers Synthesized in vitro. J. Int. Plant Biol. 2010; 52 :221–233. [ PubMed ] [ Google Scholar ]
• Cosgrove D.J. Assembly and enlargement of the primary cell wall in plants. Annu Rev Cell Dev Biol. 1997; 13 :171–201. [ PubMed ] [ Google Scholar ]
• Cosgrove D.J. Growth of the plant cell wall. Nat. Rev. Mol. Cell Biol. 2005; 6 :850–861. [ PubMed ] [ Google Scholar ]
• Crowell E.F., Gonneau M., Vernhettes S., Hofte H. Regulation of anisotropic cell expansion in higher plants. Comptes Rendus Biol. 2010a; 333 :320–324. [ PubMed ] [ Google Scholar ]
• Crowell E.F., Gonneau M., Stierhof Y.D., Hofte H., Vernhettes S. Regulated trafficking of cellulose synthases. Curr. Opin. Plant. Biol. 2010b; 13 :700–705. [ PubMed ] [ Google Scholar ]
• Crowell E.F., Bischoff V., Desprez T., Rolland A., Stierhof Y.D., Schumacher K., Gonneau M., Hofte H., Vernhettes S. Pausing of Golgi bodies on microtubules regulates secretion of cellulose synthase complexes in Arabidopsis. Plant Cell. 2009; 21 :1141–1154. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Daras G., Rigas S., Penning B., Milioni D., McCann M.C., Carpita N.C., Fasseas C., Hatzopoulos P. The thanatos mutation in Arabidopsis thaliana cellulose synthase 3 (AtCesA3) has a dominant-negative effect on cellulose synthesis and plant growth. New Phytol. 2009; 184 :114–126. [ PubMed ] [ Google Scholar ]
• Davies L.M., Harris P.J. Atomic force microscopy of microfibrils in primary cell walls. Planta. 2003; 217 :283–289. [ PubMed ] [ Google Scholar ]
• DeBolt S., Gutierrez R., Ehrhardt D.W., Melo C.V., Ross L., Cutler S.R., Somerville C., Bonetta D. Morlin, an inhibitor of cortical microtubule dynamics and cellulose synthase movement. Proc. Natl. Acad. Sci. USA. 2007; 104 :5854–5859. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Delmer D.P. CELLULOSE BIOSYNTHESIS: Exciting Times for A Difficult Field of Study. Annu Rev. Plant Physiol. Plant Mol. Biol. 1999; 50 :245–276. [ PubMed ] [ Google Scholar ]
• Desnos T., Orbovic V., Bellini C., Kronenberger J., Caboche M., Traas J., Hofte H. Procuste1 mutants identify two distinct genetic pathways controlling hypocotyl cell elongation, respectively in dark- and light-grown Arabidopsis seedlings. Development. 1996; 122 :683–693. [ PubMed ] [ Google Scholar ]
• Desprez T., Vernhettes S., Fagard M., Refregier G., Desnos T., Aletti E., Py N., Pelletier S., Hofte H. Resistance against herbicide isoxaben and cellulose deficiency caused by distinct mutations in same cellulose synthase isoform CESA6. Plant Physiol. 2002; 128 :482–490. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Desprez T., Juraniec M., Crowell E.F., Jouy H., Pochylova Z., Parcy F., Hofte H., Gonneau M., Vernhettes S. Organization of cellulose synthase complexes involved in primary cell wall synthesis in Arabidopsis thaliana. Proc. Natl. Acad. Sci. USA. 2007; 104 :15572–15577. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Dhonukshe P., Aniento F., Hwang I., Robinson D.G., Mravec J., Stierhof Y.D., Friml J. Clathrin-mediated constitutive endocytosis of PIN auxin efflux carriers in Arabidopsis. Curr. Biol. 2007; 17 :520–527. [ PubMed ] [ Google Scholar ]
• Dick-Perez M., Zhang Y., Hayes J., Salazar A., Zabotina O.A., Hong M. Structure and interactions of plant cell-wall polysaccharides by two- and three-dimensional magic-angle-spinning solid-state NMR. Biochemistry. 2011; 50 :989–1000. [ PubMed ] [ Google Scholar ]
• Doblin M.S., Kurek I., Jacob-Wilk D., Delmer D.P. Cellulose biosynthesis in plants: from genes to rosettes. Plant Cell Physiol. 2002; 43 :1407–1420. [ PubMed ] [ Google Scholar ]
• Ellis C., Turner J.G. The Arabidopsis mutant cev1 has constitutively active jasmonate and ethylene signal pathways and enhanced resistance to pathogens. Plant Cell. 2001; 13 :1025–1033. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Ellis C., Karafyllidis I., Wasternack C., Turner J.G. The Arabidopsis mutant cev1 links cell wall signaling to jasmonate and ethylene responses. Plant Cell. 2002; 14 :1557–1566. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Facette M.R., Shen Z., Bjornsdottir F.R., Briggs S.P., Smith L.G. Parallel Proteomic and Phosphoproteomic Analyses of Successive Stages of Maize Leaf Development. Plant Cell; 2013. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Fagard M., Hofte H., Vernhettes S. Cell wall mutants. Plant Physiol. Biochem. 2000a; 38 :15–25. [ Google Scholar ]
• Fagard M., Desnos T., Desprez T., Goubet F., Refregier G., Mouille G., McCann M., Rayon C., Vernhettes S., Hofte H. PROCUSTE1 encodes a cellulose synthase required for normal cell elongation specifically in roots and dark-grown hypocotyls of Arabidopsis. Plant Cell. 2000b; 12 :2409–2424. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Feraru E., Feraru M.I., Kleine-Vehn J., Martiniere A., Mouille G., Vanneste S., Vernhettes S., Runions J., Friml J. PIN polarity maintenance by the cell wall in Arabidopsis. Curr. Biol. 2011; 21 :338–343. [ PubMed ] [ Google Scholar ]
• Fernandes A.N., Thomas L.H., Altaner C.M., Callow P., Forsyth V.T., Apperley D.C., Kennedy C.J., Jarvis M.C. Nanostructure of cellulose microfibrils in spruce wood. Proc. Natl. Acad. Sci. USA. 2011; 108 :E1195–E1203. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Fisher D.D., Cyr R.J. Extending the Microtubule/Microfibril paradigm. Cellulose synthesis is required for normal cortical microtubule alignment in elongating cells. Plant Physiol. 1998; 116 :1043–1051. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Frank M., Guivarc'h A., Krupkova E., Lorenz-Meyer I., Chriqui D., Schmulling T. Tumorous shoot development (TSD) genes are required for co-ordinated plant shoot development. Plant J. 2002; 29 :73–85. [ PubMed ] [ Google Scholar ]
• Fujita M., Lechner B., Barton D.A., Overall R.L., Wasteneys G.O. The missing link: do cortical microtubules define plasma membrane nanodomains that modulate cellulose biosynthesis? Protoplasma 249 Suppl. 2012; 1 :S59–67. [ PubMed ] [ Google Scholar ]
• Fujita M., Himmelspach R., Ward J., Whittington A., Hasenbein N., Liu C., Truong T.T., Galway M.E., Mansfield S.D., Hocart C.H., Wasteneys G.O. The anisotropy1 D604N mutation in the Arabidopsis cellulose synthasel catalytic domain reduces cell wall crystallinity and the velocity of cellulose synthase complexes. Plant Physiol. 2013; 162 :74–85. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Furumizu C., Komeda Y. A novel mutation in KNOPF uncovers the role of alpha-glucosidase I during post-embryonic development in Arabidopsis thaliana. FEBS Lett. 2008; 582 :2237–2241. [ PubMed ] [ Google Scholar ]
• Gardiner J.C., Taylor N.G., Turner S.R. Control of cellulose synthase complex localization in developing xylem. Plant Cell. 2003; 15 :1740–1748. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Giddings T.H., Staehelin L.A. Microtubule-mediated control of microfibril deposition: a re-examination of the hypothesis. In: Cw L., editor. The cytoskeletal basis of plant growth and form. London: Academic; 1991. pp. 85–99. [ Google Scholar ]
• Giddings T.H., Jr, Brower D.L., Staehelin L.A. Visualization of particle complexes in the plasma membrane of Micrasterias denticulata associated with the formation of cellulose fibrils in primary and secondary cell walls. J. Cell Biol. 1980; 84 :327–339. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Gillis P.P., Mark R.E., Tang R.C. Elastic Stiffness of Crystalline Cellulose in Folded-Chain Solid State. Journal of Materials Science. 1969; 4 :1003. [ Google Scholar ]
• Gillmor C.S., Poindexter P., Lorieau J., Palcic M.M., Somerville C. Alpha-glucosidase I is required for cellulose biosynthesis and morphogenesis in Arabidopsis. J. Cell Biol. 2002; 156 :1003–1013. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Gillmor C.S., Lukowitz W., Brininstool G., Sedbrook J.C., Hamann T., Poindexter P., Somerville C. Glycosylphosphati-dylinositol-anchored proteins are required for cell wall synthesis and morphogenesis in Arabidopsis. Plant Cell. 2005; 17 :1128–1140. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Green P.B. Mechanism for Plant Cellular Morphogenesis. Science. 1962; 138 :1404–1405. [ PubMed ] [ Google Scholar ]
• Gu Y., Somerville C. Cellulose synthase interacting protein: a new factor in cellulose synthesis. Plant Signal. Behav. 2010a; 5 :1571–1574. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Gu Y., Kaplinsky N., Bringmann M., Cobb A., Carroll A., Sampath-kumar A., Baskin T.I., Persson S., Somerville C.R. Identification of a cellulose synthase-associated protein required for cellulose biosynthesis. Proc. Natl. Acad. Sci. USA. 2010b; 107 :12866–12871. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Guerriero G., Fugelstad J., Bulone V. What Do We Really Know about Cellulose Biosynthesis in Higher Plants? J. Integr. Plant Biol. 2010; 52 :161–175. [ PubMed ] [ Google Scholar ]
• Gutierrez R., Lindeboom J.J., Paredez A.R., Emons A.M., Ehrhardt D.W. Arabidopsis cortical microtubules position cellulose synthase delivery to the plasma membrane and interact with cellulose synthase trafficking compartments. Nat. Cell Biol. 2009; 11 :797–806. [ PubMed ] [ Google Scholar ]
• Ha M.A., Apperley D.C., Evans B.W., Huxham M., Jardine W.G., Vietor R.J., Reis D., Vian B., Jarvis M.C. Fine structure in cellulose microfibrils: NMR evidence from onion and quince. Plant J. 1998; 16 :183–190. [ PubMed ] [ Google Scholar ]
• Haigler C.H., Brown R.M. Transport of Rosettes from the Golgi-Apparatus to the Plasma-Membrane in Isolated Mesophyll-Cells of Zinnia-Elegans during Differentiation to Tracheary Elements in Suspension-Culture. Protoplasma. 1986; 134 :111–120. [ Google Scholar ]
• Haigler C.H., Malcolmbrown R., Benziman M. Calcofluor White St Alters the Invivo Assembly of Cellulose Microfibrils. Science. 1980; 210 :903–906. [ PubMed ] [ Google Scholar ]
• Hamann T., Osborne E., Youngs H.L., Misson J., Nussaume L., Somerville C. Global expression analysis of CESA and CSL genes in Arabidopsis. Cellulose. 2004; 11 :279–286. [ Google Scholar ]
• Hamant O., Heisler M.G., Jonsson H., Krupinski P., Uyttewaal M., Bokov P., Corson F., Sahlin P., Boudaoud A., Meyerowitz E.M., Couder Y., Traas J. Developmental patterning by mechanical signals in Arabidopsis. Science. 2008; 322 :1650–1655. [ PubMed ] [ Google Scholar ]
• Harpaz-Saad S., Western T.L., Kieber J.J. The FEI2-SOS5 pathway and CELLULOSE SYNTHASE 5 are required for cellulose biosynthesis in the Arabidopsis seed coat and affect pectin mucilage structure. Plant Signal. Behav. 2012; 7 :285–288. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Harpaz-Saad S., McFarlane H.E., Xu S., Divi U.K., Forward B., Western T.L., Kieber J.J. Cellulose synthesis via the FEI2 RLK/SOS5 pathway and cellulose synthase 5 is required for the structure of seed coat mucilage in Arabidopsis. Plant J. 2011; 68 :941–953. [ PubMed ] [ Google Scholar ]
• Harris D., Bulone V., Ding S.Y., DeBolt S. Tools for cellulose analysis in plant cell walls. Plant Physiol. 2010; 153 :420–426. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Harris D.M., Corbin K., Wang T., Gutierrez R., Bertolo A.L., Petti C., Smilgies D.M., Estevez J.M., Bonetta D., Urbanowicz B.R., Ehrhardt D.W., Somerville C.R., Rose J.K., Hong M., Debolt S. Cellulose microfibril crystallinity is reduced by mutating C-terminal transmembrane region residues CESA1A903V and CESA3T942I of cellulose synthase. Proc. Natl. Acad. Sci. USA. 2012; 109 :4098–4103. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Hauser M.T., Morikami A., Benfey P.N. Conditional root expansion mutants of Arabidopsis. Development. 1995; 121 :1237–1252. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Hayashi T., Yoshida K., Park Y.W., Konishi T., Baba K. Cellulose metabolism in plants. Intl. Rev. Cytol. 2005; 247 :1–34. [ PubMed ] [ Google Scholar ]
• Heath I.B. A unified hypothesis for the role of membrane bound enzyme complexes and microtubules in plant cell wall synthesis. J Theor. Biol. 1974; 48 :445–449. [ PubMed ] [ Google Scholar ]
• Hematy K., Sado P.E., Van Tuinen A., Rochange S., Desnos T., Balzergue S., Pelletier S., Renou J.P., Hofte H. A receptor-like kinase mediates the response of Arabidopsis cells to the inhibition of cellulose synthesis. Curr. Biol. 2007; 17 :922–931. [ PubMed ] [ Google Scholar ]
• Hepler P.K., Newcomb E.H. Microtubules and Fibrils in the Cytoplasm of Coleus Cells Undergoing Secondary Wall Deposition. J Cell. Biol. 1964; 20 :529–532. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Hermans C., Porco S., Verbruggen N., Bush D.R. Chitinase-like protein CTL1 plays a role in altering root system architecture in response to multiple environmental conditions. Plant Physiol. 2010; 152 :904–917. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Hernandez-Bianco C., Feng D.X., Hu J., Sanchez-Vallet A., Deslandes L., Llorente F., Berrocal-Lobo M., Keller H., Bariet X., Sanchez-Rodriguez C., Anderson L.K., Somerville S., Marco Y., Molina A. Impairment of cellulose synthases required for Arabidopsis secondary cell wall formation enhances disease resistance. Plant Cell. 2007; 19 :890–903. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Hieu H.C., Tuan N.A., Li H.Y., Miyauchi Y., Mizutani G. Sum Frequency Generation Microscopy Study of Cellulose Fibers. Applied Spectro. 2011; 65 :1254–1259. [ PubMed ] [ Google Scholar ]
• Himmelspach R., Williamson R.E., Wasteneys G.O. Cellulose microfibril alignment recovers from DCB-induced disruption despite microtubule disorganization. Plant J. 2003; 36 :565–575. [ PubMed ] [ Google Scholar ]
• His I., Driouich A., Nicol F., Jauneau A., Hofte H. Altered pectin composition in primary cell walls of korrigan, a dwarf mutant of Arabidopsis deficient in a membrane-bound endo-1,4-beta-glucanase. Planta. 2001; 212 :348–358. [ PubMed ] [ Google Scholar ]
• Jacob-Wilk D., Kurek I., Hogan P., Delmer D.P. The cotton fiber zinc-binding domain of cellulose synthase A1 from Gossypium hirsutum displays rapid turnover in vitro and in vivo. Proc. Natl. Acad. Sci. USA. 2006; 103 :12191–12196. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Kakar K., Zhang H., Scheres B., Dhonukshe P. CLASP-mediated cortical microtubule organization guides PIN polarization axis. Nature. 2013; 495 :529–533. [ PubMed ] [ Google Scholar ]
• Keegstra K. Plant Cell Walls. Plant Physiol. 2010; 154 :483–486. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Keegstra K., Talmadge K.W., Bauer W.D., Albersheim P. The Structure of Plant Cell Walls: III. A Model of the Walls of Suspension-cultured Sycamore Cells Based on the Interconnections of the Macromolecular Components. Plant Physiol. 1973; 51 :188–197. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Kerstens S., Verbelen J.P. Cellulose orientation in the outer epidermal wall of angiosperm roots: implications for biosystematics. Annals Bot. 2002; 90 :669–676. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Kim W.C., Ko J.H., Han K.H. Identification of a cis-acting regulatory motif recognized by MYB46, a master transcriptional regulator of secondary wall biosynthesis. Plant Mol. Biol. 2012a; 78 :489–501. [ PubMed ] [ Google Scholar ]
• Kim W.C., Kim J.Y., Ko J.H., Kim J., Han K.H. Transcription factor MYB46 is an obligate component of the transcriptional regulatory complex for functional expression of secondary wall-associated cellulose synthases in Arabidopsis thaliana. Plant Physiol. 2013; 170 :1374–1378. [ PubMed ] [ Google Scholar ]
• Kim W.C., Ko J.H., Kim J.Y., Kim J.M., Bae H.J., Han K.H. Plant J. 2012b. MYB46 directly regulates the gene expression of secondary wall-associated cellulose synthases in Arabidopsis. [ PubMed ] [ Google Scholar ]
• Kimura S., Laosinchai W., Itoh T., Cui X., Linder C.R., Brown R.M., Jr Immunogold labeling of rosette terminal cellulose-synthesizing complexes in the vascular plant vigna angularis. Plant Cell. 1999; 11 :2075–2086. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Ko J.H., Kim W.C., Han K.H. Ectopic expression of MYB46 identifies transcriptional regulatory genes involved in secondary wall biosynthesis in Arabidopsis. Plant J. 2009; 60 :649–665. [ PubMed ] [ Google Scholar ]
• Ko J.H., Kim J.H., Jayanty S.S., Howe G.A., Han K.H. Loss of function of COBRA, a determinant of oriented cell expansion, invokes cellular defence responses in Arabidopsis thaliana. J. Exp. Bot. 2006; 57 :2923–2936. [ PubMed ] [ Google Scholar ]
• Koyama M., Helbert W., Imai T., Sugiyama J., Henrissat B. Parallel-up structure evidences the molecular directionality during biosynthesis of bacterial cellulose. Proc. Natl. Acad. Sci. USA. 1997; 94 :9091–9095. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Krupkova E., Schmulling T. Developmental consequences of the tumorous shoot development1 mutation, a novel allele of the cellulose-synthesizing KORRIGAN1 gene. Plant Mol Biol. 2009; 71 :641–655. [ PubMed ] [ Google Scholar ]
• Kudlicka K., Brown Jr R.M. Cellulose and Callose Biosynthesis in Higher Plants (I. Solubilization and Separation of (1->3)-and (1->4)-[beta]-Glucan Synthase Activities from Mung Bean). Plant Physiol. 1997; 115 :643–656. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Kudlicka K., Brown R.M., Jr, Li L., Lee J.H., Shin H., Kuga S. [beta]-Glucan Synthesis in the Cotton Fiber (IV. In Vitro Assembly of the Cellulose I Allomorph). Plant Physiol. 1995; 107 :111–123. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Kuga S., Brown R.M. Silver labeling of the reducing ends of bacterial cellulose. Carbo. Research. 1988; 180 :345–350. [ Google Scholar ]
• Kurek I., Kawagoe Y., Jacob-Wilk D., Doblin M., Delmer D. Dimerization of cotton fiber cellulose synthase catalytic subunits occurs via oxidation of the zinc-binding domains. Proc. Natl. Acad. Sci. USA. 2002; 99 :11109–11114. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Kwon Y., Kim S.H., Jung M.S., Kim M.S., Oh J.E., Ju H.W., Kim K.I., Vierling E., Lee H., Hong S.W. Arabidopsis hot2 encodes an endochitinase-like protein that is essential for tolerance to heat, salt and drought stresses. Plant J. 2007; 49 :184–193. [ PubMed ] [ Google Scholar ]
• L. Segal J.J.C., Martin A. E., Conrad C. M. An Empirical Method for Estimating the Degree of Crystallinity of Native Cellulose Using the X-Ray Diffractometer. Text Res. J. 1959; 29 :786–794. [ Google Scholar ]
• Lai-Kee-Him J., Chanzy H., Muller M., Putaux J.L., Imai T., Bulone V. In vitro versus in vivo cellulose microfibrils from plant primary wall synthases: structural differences. J. Biol. Chem. 2002; 277 :36931–36939. [ PubMed ] [ Google Scholar ]
• Lane D.R., Wiedemeier A., Peng L., Hofte H., Vernhettes S., Desprez T., Hocart C.H., Birch R.J., Baskin T.I., Burn J.E., Arioli T., Betzner A.S., Williamson R.E. Temperature-sensitive alleles of RSW2 link the KORRIGAN endo-1,4-beta-glucanase to cellulose synthesis and cytokinesis in Arabidopsis. Plant Physiol. 2001; 126 :278–288. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Ledbetter M.C., Porter K.R. A “Microtubule” in Plant Cell Fine Structure. J. Cell Biol. 1963; 19 :239–250. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Lei L., Li S., Gu Y. Cellulose synthase interactive protein 1 (CSI1) mediates the intimate relationship between cellulose microfibrils and cortical microtubules. Plant Signal. Behav. 2012a; 7 :714–718. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Lei L., Li S., Gu Y. Cellulose synthase complexes: composition and regulation. Front. Plant Sci. 2012b; 3 :75. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Lertpiriyapong K., Sung Z.R. The elongation defective1 mutant of Arabidopsis is impaired in the gene encoding a serine-rich secreted protein. Plant Mol. Biol. 2003; 53 :581–595. [ PubMed ] [ Google Scholar ]
• Levy I., Shani Z., Shoseyov O. Modification of polysaccharides and plant cell wall by endo-1,4-beta-glucanase and cellulose-binding domains. Biomol. Engineer. 2002; 19 :17–30. [ PubMed ] [ Google Scholar ]
• Li R., Liu P., Wan Y., Chen T., Wang Q., Mettbach U., Baluska F., Samaj J., Fang X., Lucas W.J., Lin J. A membrane microdomain-associated protein, Arabidopsis Flot1, is involved in a clathrin-independent endocytic pathway and is required for seedling development. Plant Cell. 2012a; 24 :2105–2122. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Li S., Gu Y. Cellulose biosynthesis in higher plants and the role of the cytoskeleton. In: eLS A.M. Hetherington., editor. Chichester: John Wiley and Sons; 2012b. [ Google Scholar ]
• Li S., Lei L., Gu Y. Functional analysis of complexes with mixed primary and secondary cellulose synthases. Plant Signal. Behav. 2013:8. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Li S., Lei L., Somerville C., Gu Y. Cellulose synthase interacting protein 1 (CSI1) links microtubules and cellulose synthase complexes. Proc. Natl. Acad. Sci. USA. 2012c; 109 :185–190. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Liang Y.K., Xie X., Lindsay S.E., Wang Y.B., Masle J., Williamson L., Leyser O., Hetherington A.M. Cell wall composition contributes to the control of transpiration efficiency in Arabidopsis thaliana. Plant J. 2010; 64 :679–686. [ PubMed ] [ Google Scholar ]
• Liebminger E., Grass J., Altmann F., Mach L., Strasser R. Characterizing the Link between Glycosylation State and Enzymatic Activity of the Endo-beta1,4-glucanase KORRIGAN1 from Arabidopsis thaliana. J. Biol. Chem. 2013; 288 :22270–22280. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Lin D., Cao L., Zhou Z., Zhu L., Ehrhardt D., Yang Z., Fu Y. Rho GTPase signaling activates microtubule severing to promote microtubule ordering in Arabidopsis. Curr Biol. 2013; 23 :290–297. [ PubMed ] [ Google Scholar ]
• Lloyd C., Chan J. The parallel lives of microtubules and cellulose microfibrils. Curr. Opin. Plant Biol. 2008; 11 :641–646. [ PubMed ] [ Google Scholar ]
• Lukowitz W., Nickle T.C., Meinke D.W., Last R.L., Conklin P.L., Somerville C.R. Arabidopsis cytl mutants are deficient in a mannose-1 -phosphate guanylyltransferase and point to a requirement of N-linked glycosylation for cellulose biosynthesis. Proc. Natl. Acad. Sci. USA. 2001; 98 :2262–2267. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Master E.R., Rudsander U.J., Zhou W., Henriksson H., Divne C., Denman S., Wilson D.B., Teeri T.T. Recombinant expression and enzymatic characterization of PttCel9A, a KOR homologue from Populus tremula x tremuloides. Biochemistry. 2004; 43 :10080–10089. [ PubMed ] [ Google Scholar ]
• Maurer A., Fengel D. Parallel Orientation of the Molecular Chains in Cellulose-I and Cellulose-Ii Deriving from Higher-Plants. Holz Als Roh-Und Werkstoff. 1992; 50 :493–493. [ Google Scholar ]
• McCarthy R.L., Zhong R., Ye Z.H. MYB83 is a direct target of SND1 and acts redundantly with MYB46 in the regulation of secondary cell wall biosynthesis in Arabidopsis. Plant Cell Physiol. 2009; 50 :1950–1964. [ PubMed ] [ Google Scholar ]
• Mendu V., Stork J., Harris D., DeBolt S. Cellulose synthesis in two secondary cell wall processes in a single cell type. Plant Signal. Behav. 2011a; 6 :1638–1643. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Mendu V., Griffiths J.S., Persson S., Stork J., Downie A.B., Voiniciuc C., Haughn G.W., DeBolt S. Subfunctionalization of Cellulose Synthases in Seed Coat Epidermal Cells Mediates Secondary Radial Wall Synthesis and Mucilage Attachment. Plant Physiol. 2011b; 157 :441–453. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Mitsuda N., Iwase A., Yamamoto H., Yoshida M., Seki M., Shinozaki K., Ohme-Takagi M. NAC transcription factors, NST1 and NST3, are key regulators of the formation of secondary walls in woody tissues of Arabidopsis. Plant Cell. 2007; 19 :270–280. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Molhoj M., Ulvskov P., Dal Degan F. Characterization of a functional soluble form of a Brassica napus membrane-anchored endo-1,4-beta-glucanase heterologously expressed in Pichia pastoris. Plant Physiol. 2001; 127 :674–684. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Molhoj M., Pagant S., Hofte H. Towards understanding the role of membrane-bound endo-beta-1,4-glucanases in cellulose biosynthesis. Plant Cell Physiol. 2002; 43 :1399–1406. [ PubMed ] [ Google Scholar ]
• Montezinos D., Brown M., Jr Surface architecture of the plant cell: biogenesis of the cell wall, with special emphasis on the role of the plasma membrane in cellulose biosynthesis. J. Supramol. Struct. 1976; 5 :277–290. [ PubMed ] [ Google Scholar ]
• Morgan J.L., Strumillo J., Zimmer J. Crystallographic snapshot of cellulose synthesis and membrane translocation. Nature. 2013; 493 :181–186. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Mueller S.C., Brown R.M., Jr Evidence for an intramem-brane component associated with a cellulose microfibril-synthesizing complex in higher plants. J. Cell Biol. 1980; 84 :315–326. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Nakamura M., Hashimoto T. A mutation in the Arabidopsis gamma-tubulin-containing complex causes helical growth and abnormal microtubule branching. J Cell Sci. 2009; 122 :2208–2217. [ PubMed ] [ Google Scholar ]
• Newman R.H., Hill S.J., Harris P.J. Wide-angle x-ray scattering and solid-state nuclear magnetic resonance data combined to test models for cellulose microfibrils in mung bean cell walls. Plant Physiol. 2013; 163 :1558–1567. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Nickle T.C., Meinke D.W. A cytokinesis-defective mutant of Arabidopsis (cyt1) characterized by embryonic lethality, incomplete cell walls, and excessive callose accumulation. Plant J. 1998; 15 :321–332. [ PubMed ] [ Google Scholar ]
• Nicol F., His I., Jauneau A., Vernhettes S., Canut H., Hofte H. A plasma membrane-bound putative endo-1,4-beta-D-glucanase is required for normal wall assembly and cell elongation in Arabidopsis. EMBO J. 1998; 17 :5563–5576. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Nuhse T.S., Stensballe A., Jensen O.N., Peck S.C. Phosphoproteomics of the Arabidopsis plasma membrane and a new phosphorylation site database. Plant Cell. 2004; 16 :2394–2405. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Ohlsson A.B., Djerbi S., Winzell A., Bessueille L., Staldal V., Li X.G., Blomqvist K., Bulone V., Teeri T.T., Berglund T. Cell suspension cultures of Populus tremula x P-tremuloides exhibit a high level of cellulose synthase gene expression that coincides with increased in vitro cellulose synthase activity. Protoplasma. 2006; 228 :221–229. [ PubMed ] [ Google Scholar ]
• Okuda K., Li L., Kudlicka K., Kuga S., Brown R.M., Jr [beta]-Glucan Synthesis in the Cotton Fiber (I. Identification of [beta]-1,4-and [beta]-1,3-Glucans Synthesized in Vitro). Plant Physiol. 1993; 101 :1131–1142. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Pagant S., Bichet A., Sugimoto K., Lerouxel O., Desprez T., Mc-Cann M., Lerouge P., Vernhettes S., Hofte H. KOBITO1 encodes a novel plasma membrane protein necessary for normal synthesis of cellulose during cell expansion in Arabidopsis. Plant Cell. 2002; 14 :2001–2013. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Paradez A., Wright A., Ehrhardt D.W. Microtubule cortical array organization and plant cell morphogenesis. Curr. Opin. Plant Biol. 2006; 9 :571–578. [ PubMed ] [ Google Scholar ]
• Paredez A.R., Somerville C.R., Ehrhardt D.W. Visualization of cellulose synthase demonstrates functional association with microtubules. Science. 2006; 312 :1491–1495. [ PubMed ] [ Google Scholar ]
• Paredez A.R., Persson S., Ehrhardt D.W., Somerville C.R. Genetic evidence that cellulose synthase activity influences microtubule cortical array organization. Plant Physiol. 2008; 147 :1723–1734. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Park S., Baker J.O., Himmel M.E., Parilla P.A., Johnson D.K. Cellulose crystallinity index: measurement techniques and their impact on interpreting cellulase performance. Biotechnol. Biofuels. 2010; 3 :10. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Park Y.B., Lee C.M., Koo B.W., Park S., Cosgrove D.J., Kim S.H. Monitoring meso-scale ordering of cellulose in intact plant cell walls using sum frequency generation spectroscopy. Plant Physiol. 2013; 163 :907–913. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Pauly M., Keegstra K. Plant cell wall polymers as precursors for biofuels. Curr. Opin. Plant Biol. 2010; 13 :305–312. [ PubMed ] [ Google Scholar ]
• Pear J.R., Kawagoe Y., Schreckengost W.E., Delmer D.P., Stalker D.M. Higher plants contain homologs of the bacterial celA genes encoding the catalytic subunit of cellulose synthase. Proc. Natl. Acad. Sci. USA. 1996; 93 :12637–12642. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Peng L.C., Kawagoe Y., Hogan P., Delmer D. Sitosterol-beta-glucoside as primer for cellulose synthesis in plants. Science. 2002; 295 :147–150. [ PubMed ] [ Google Scholar ]
• Persson S., Wei H., Milne J., Page G.P., Somerville C.R. Identification of genes required for cellulose synthesis by regression analysis of public microarray data sets. Proc. Natl. Acad. Sci. USA. 2005; 102 :8633–8638. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Persson S., Paredez A., Carroll A., Palsdottir H., Doblin M., Poindexter P., Khitrov N., Auer M., Somerville C.R. Genetic evidence for three unique components in primary cell-wall cellulose synthase complexes in Arabidopsis. Proc. Natl. Acad. Sci. USA. 2007; 104 :15566–15571. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Plomion C., Leprovost G., Stokes A. Wood formation in trees. Plant Physiol. 2001; 127 :1513–1523. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Popper Z.A. Evolution and diversity of green plant cell walls. Curr. Opin. Plant Biol. 2008; 11 :286–292. [ PubMed ] [ Google Scholar ]
• Pysh L., Alexander N., Swatzyna L., Harbert R. Four alleles of AtCESA3 form an allelic series with respect to root phenotype in Arabidopsis thaliana. Physiol. Planta. 2012; 144 :369–381. [ PubMed ] [ Google Scholar ]
• Reiter W.D., Chapple C., Somerville C.R. Mutants of Arabidopsis thaliana with altered cell wall polysaccharide composition. Plant J. 1997; 12 :335–345. [ PubMed ] [ Google Scholar ]
• Richmond T. Higher plant cellulose synthases. Genome Biol. 2000; 1 :REVIEWS3001. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Richmond T.A., Somerville C.R. The cellulose synthase superfamily. Plant Physiol. 2000; 124 :495–498. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Robert S., Bichet A., Grandjean O., Kierzkowski D., Satiat-Jeunemaitre B., Pelletier S., Hauser M.T., Hofte H., Vernhettes S. An Arabidopsis endo-1,4-beta-D-glucanase involved in cellulose synthesis undergoes regulated intracellular cycling. Plant Cell. 2005; 17 :3378–3389. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Roberts A., Roberts E. Evolution of the cellulose synthase (CesA) gene family: insights from green algae and seedless plants. In: Brown R.M., Saxena I.M., editors. Cellulose: Molecular and Structural Biology. Dordrecht: Springer; 2007. pp. 17–34. In. [ Google Scholar ]
• Roberts K., McCann M.C. Xylogenesis: the birth of a corpse. Curr. Opin. Plant Biol. 2000; 3 :517–522. [ PubMed ] [ Google Scholar ]
• Rogers L.A., Dubos C., Surman C., Willment J., Cullis I.F., Mansfield S.D., Campbell M.M. Comparison of lignin deposition in three ectopic lignification mutants. New Phytol. 2005; 168 :123–140. [ PubMed ] [ Google Scholar ]
• Roland J.C., Vian B., Reis D. Observations with cytochemistry and ultracryotomy on the fine structure of the expanding walls in actively elongating plant cells. J. Cell Sci. 1975; 19 :239–259. [ PubMed ] [ Google Scholar ]
• Ross P., Mayer R., Benziman M. Cellulose biosynthesis and function in bacteria. Microbiol. Rev. 1991; 55 :35–58. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Roudier F., Fernandez A.G., Fujita M., Himmelspach R., Borner G.H., Schindelman G., Song S., Baskin T.I., Dupree P., Wasteneys G.O., Benfey P.N. COBRA, an Arabidopsis extracellular glycosyl-phosphatidyl inositol-anchored protein, specifically controls highly anisotropic expansion through its involvement in cellulose microfibril orientation. Plant Cell. 2005; 17 :1749–1763. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Rubio-Diaz S., Perez-Perez J.M., Gonzalez-Bayon R., Munoz-Viana R., Borrega N., Mouille G., Hernandez-Romero D., Robles P., Hofte H., Ponce M.R., Micol J.L. Cell Expansion-Mediated Organ Growth Is Affected by Mutations in Three EXIGUA Genes. PloS one. 2012:7. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Rudsander U.J., Sandstrom C., Piens K., Master E.R., Wilson D.B., Brumer Iii H., Kenne L., Teeri T.T. Comparative NMR analysis of cellooligosaccharide hydrolysis by GH9 bacterial and plant endo-1,4-beta-glucanases. Biochemistry. 2008; 47 :5235–5241. [ PubMed ] [ Google Scholar ]
• Sanchez-Rodriguez C., Bauer S., Hematy K., Saxe F., Ibanez A.B., Vodermaier V., Konlechner C., Sampathkumar A., Ruggeberg M., Aichinger E., Neumetzier L., Burgert I., Somerville C., Hauser M.T., Persson S. Chitinase-like1/pom-pom1 and its homolog CTL2 are glucan-interacting proteins important for cellulose biosynthesis in Arabidopsis. Plant Cell. 2012; 24 :589–607. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Sato S., Kato T., Kakegawa K., Ishii T., Liu Y.G., Awano T., Takabe K., Nishiyama Y., Kuga S., Nakamura Y., Tabata S., Shibata D. Role of the putative membrane-bound endo-1,4-beta-glucanase KORRIGAN in cell elongation and cellulose synthesis in Arabidopsis thaliana. Plant Cell Physiol. 2001; 42 :251–263. [ PubMed ] [ Google Scholar ]
• Saxena I.M., Brown R.M., Jr Cellulose biosynthesis: current views and evolving concepts. Ann. Bot. 2005; 96 :9–21. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Saxena I.M., Brown R.M., Dandekar T. Structure-function characterization of cellulose synthase: relationship to other glycosyl-transferases. Phytochemistry. 2001; 57 :1135–1148. [ PubMed ] [ Google Scholar ]
• Scheible W.R., Eshed R., Richmond T., Delmer D., Somerville C. Modifications of cellulose synthase confer resistance to isoxaben and thiazolidinone herbicides in Arabidopsis Ixr1 mutants. Proc. Natl. Acad. Sci. USA. 2001; 98 :10079–10084. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Scheible W.R., Fry B., Kochevenko A., Schindelasch D., Zimmerli L., Somerville S., Loria R., Somerville C.R. An Arabidopsis mutant resistant to thaxtomin A, a cellulose synthesis inhibitor from Streptomyces species. Plant Cell. 2003; 15 :1781–1794. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Scheller H.V., Ulvskov P. Hemicelluloses. Annu. Rev. Plant Biol. 2010; 61 :263–289. [ PubMed ] [ Google Scholar ]
• Schindelman G., Morikami A., Jung J., Baskin T.I., Carpita N.C., Derbyshire P., McCann M.C., Benfey P.N. COBRA encodes a putative GPI-anchored protein, which is polarly localized and necessary for oriented cell expansion in Arabidopsis. Genes Dev. 2001; 15 :1115–1127. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Schrick K. A link between sterol biosynthesis, the cell-wall, and cellulose in Arabidopsis (vol 38, pg 227, 2004). Plant J. 2004; 38 :562–562. [ PubMed ] [ Google Scholar ]
• Schrick K., Debolt S., Bulone V. Deciphering the molecular functions of sterols in cellulose biosynthesis. Front. Plant Sci. 2012; 3 :84. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Sethaphong L., Haigler C.H., Kubicki J.D., Zimmer J., Bonetta D., DeBolt S., Yingling Y.G. Tertiary model of a plant cellulose synthase. Proc. Natl. Acad. Sci. USA. 2013; 110 :7512–7517. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Slabaugh E., Davis J.K., Haigler C.H., Yingling Y.G., Zimmer J. Cellulose synthases: new insights from crystallography and modeling. Trends Plant Sci; 2013. [ PubMed ] [ Google Scholar ]
• Somerville C. Cellulose synthesis in higher plants. Annu. Rev. Cell Dev. Biol. 2006; 22 :53–78. [ PubMed ] [ Google Scholar ]
• Somerville C. Biofuels. Curr. Biol. 2007; 17 :R115–R119. [ PubMed ] [ Google Scholar ]
• Stork J., Harris D., Griffiths J., Williams B., Beisson F., Li-Beisson Y., Mendu V., Haughn G., DeBolt S. CELLULOSE SYNTHASE9 Serves a Nonredundant Role in Secondary Cell Wall Synthesis in Arabidopsis Epidermal Testa Cells. Plant Physiol. 2010; 153 :580–589. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Sturcova A., His I., Apperley D.C., Sugiyama J., Jarvis M.C. Structural details of crystalline cellulose from higher plants. Biomacromolecules. 2004; 5 :1333–1339. [ PubMed ] [ Google Scholar ]
• Sugimoto K., Himmelspach R., Williamson R.E., Wasteneys G.O. Mutation or drug-dependent microtubule disruption causes radial swelling without altering parallel cellulose microfibril deposition in Arabidopsis root cells. Plant Cell. 2003; 15 :1414–1429. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Szyjanowicz P.M., McKinnon I., Taylor N.G., Gardiner J., Jarvis M.C., Turner S.R. The irregular xylem 2 mutant is an allele of korrigan that affects the secondary cell wall of Arabidopsis thaliana. Plant J. 2004; 37 :730–740. [ PubMed ] [ Google Scholar ]
• Taylor N.G. Identification of cellulose synthase AtCesA7 (IRX3) in vivo phosphorylation sites-a potential role in regulating protein degradation. Plant Mol. Biol. 2007; 64 :161–171. [ PubMed ] [ Google Scholar ]
• Taylor N.G., Laurie S., Turner S.R. Multiple cellulose synthase catalytic subunits are required for cellulose synthesis in Arabidopsis. Plant Cell. 2000; 12 :2529–2540. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Taylor N.G., Scheible W.R., Cutler S., Somerville C.R., Turner S.R. The irregular xylem3 locus of Arabidopsis encodes a cellulose synthase required for secondary cell wall synthesis. Plant Cell. 1999; 11 :769–780. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Taylor N.G., Howells R.M., Huttly A.K., Vickers K., Turner S.R. Interactions among three distinct CesA proteins essential for cellulose synthesis. Proc. Natl. Acad. Sci. USA. 2003; 100 :1450–1455. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Thomas L.H., Forsyth V.T., Sturcova A., Kennedy C.J., May R.P., Altaner C.M., Apperley D.C., Wess T.J., Jarvis M.C. Structure of cellulose microfibrils in primary cell walls from collenchyma. Plant Physiol. 2013; 161 :465–476. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Timmers J., Vernhettes S., Desprez T., Vincken J.P., Visser R.G., Trindade L.M. Interactions between membrane-bound cellulose synthases involved in the synthesis of the secondary cell wall. FEBS letters. 2009; 583 :978–982. [ PubMed ] [ Google Scholar ]
• Tsekos I. The sites of cellulose synthesis in algae: Diversity and evolution of cellulose-synthesizing enzyme complexes. J. Phycol. 1999; 35 :635–655. [ Google Scholar ]
• Turner S.R., Somerville C.R. Collapsed xylem phenotype of Arabidopsis identifies mutants deficient in cellulose deposition in the secondary cell wall. Plant Cell. 1997; 9 :689–701. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Wang J., Howles P.A., Cork A.H., Birch R.J., Williamson R.E. Chimeric proteins suggest that the catalytic and/or C-terminal domains give CesA1 and CesA3 access to their specific sites in the cellulose synthase of primary walls. Plant Physiol. 2006; 142 :685–695. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Wang T., Zabotina O., Hong M. Pectin-cellulose interactions in the Arabidopsis primary cell wall from two-dimensional magic-angle-spinning solid-state nuclear magnetic resonance. Biochemistry. 2012; 51 :9846–9856. [ PubMed ] [ Google Scholar ]
• Wasteneys G.O. Progress in understanding the role of microtubules in plant cells. Curr. Opin. Plant Biol. 2004; 7 :651–660. [ PubMed ] [ Google Scholar ]
• Wasteneys G.O., Collings D.A. Expanding beyong the great divide: the cytoskeleton and axial growth. In: Hussey P.J., editor. The plant sytoskeleton in cell differentiation and development. Blackwell; 2004. pp. 83–115. [ Google Scholar ]
• Webb M., Jouannic S., Foreman J., Linstead P., Dolan L. Cell specification in the Arabidopsis root epidermis requires the activity of ECTOPIC ROOT HAIR 3--a katanin-p60 protein. Development. 2002; 129 :123–131. [ PubMed ] [ Google Scholar ]
• Wightman R., Turner S.R. The roles of the cytoskeleton during cellulose deposition at the secondary cell wall. Plant J. 2008; 54 :794–805. [ PubMed ] [ Google Scholar ]
• Wightman R., Turner S. Trafficking of the plant cellulose synthase complex. Plant Physiol. 2010a; 153 :427–432. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Wightman R., Turner S. Trafficking of the cellulose synthase complex in developing xylem vessels. Biochemical Society Transactions. 2010b; 38 :755–760. [ PubMed ] [ Google Scholar ]
• Wightman R., Marshall R., Turner S.R. A cellulose synthase-containing compartment moves rapidly beneath sites of secondary wall synthesis. Plant Cell Physiol. 2009; 50 :584–594. [ PubMed ] [ Google Scholar ]
• Williamson R.E., Burn J.E., Hocart C.H. Towards the mechanism of cellulose synthesis. Trends Plant Sci. 2002; 7 :461–467. [ PubMed ] [ Google Scholar ]
• Wolf S., Hematy K., Hofte H. Growth control and cell wall signaling in plants. Ann. Rev. Plant Biol. 2012; 63 :381–407. [ PubMed ] [ Google Scholar ]
• Xie L., Yang C., Wang X. Brassinosteroids can regulate cellulose biosynthesis by controlling the expression of CESA genes in Arabidopsis. J. Exp. Bot. 2011; 62 :4495–4506. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Yamaoka S., Shimono Y., Shirakawa M., Fukao Y., Kawase T., Hatsugai N., Tamura K., Shimada T., Hara-Nishimura I. Identification and dynamics of Arabidopsis adaptor protein-2 complex and its involvement in floral organ development. Plant Cell. 2013; 25 :2958–2969. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Zhang T.M.-L., S. Tittmann B., Cosgrove DJ. Visualization of the nanoscale pattern of recently-deposited cellulose microfibrils and matrix materials in never-dried primary walls of the onion epidermis. 2013. Cellulose DOI 10.1007/s10570-013-9996-1.
• Zhong R., Ye Z.H. MYB46 and MYB83 bind to the SMRE sites and directly activate a suite of transcription factors and secondary wall biosynthetic genes. Plant Cell Physiol. 2012; 53 :368–380. [ PubMed ] [ Google Scholar ]
• Zhong R., Demura T., Ye Z.H. SND1, a NAC domain transcription factor, is a key regulator of secondary wall synthesis in fibers of Arabidopsis. Plant Cell. 2006; 18 :3158–3170. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Zhong R., Richardson E.A., Ye Z.H. Two NAC domain transcription factors, SND1 and NST1, function redundantly in regulation of secondary wall synthesis in fibers of Arabidopsis. Planta. 2007a; 225 :1603–1611. [ PubMed ] [ Google Scholar ]
• Zhong R., Richardson E.A., Ye Z.H. The MYB46 transcription factor is a direct target of SND1 and regulates secondary wall biosynthesis in Arabidopsis. Plant Cell. 2007b; 19 :2776–2792. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Zhong R., Lee C., Ye Z.H. Evolutionary conservation of the transcriptional network regulating secondary cell wall biosynthesis. Trends Plant Sci. 2010; 15 :625–632. [ PubMed ] [ Google Scholar ]
• Zhong R., Kays S.J., Schroeder B.P., Ye Z.H. Mutation of a chitinase-like gene causes ectopic deposition of lignin, aberrant cell shapes, and overproduction of ethylene. Plant Cell. 2002; 14 :165–179. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Zhong R., Morrison W.H., 3rd, Freshour G.D., Hahn M.G., Ye Z.H. Expression of a mutant form of cellulose synthase AtCesA7 causes dominant negative effect on cellulose biosynthesis. Plant Physiol. 2003; 132 :786–795. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Zhong R., Lee C., Zhou J., McCarthy R.L., Ye Z.H. A battery of transcription factors involved in the regulation of secondary cell wall biosynthesis in Arabidopsis. Plant Cell. 2008; 20 :2763–2782. [ PMC free article ] [ PubMed ] [ Google Scholar ]
• Zuo J., Niu Q.W., Nishizawa N., Wu Y., Kost B., Chua N.H. KORRIGAN, an Arabidopsis endo-1,4-beta-glucanase, localizes to the cell plate by polarized targeting and is essential for cytokinesis. Plant Cell. 2000; 12 :1137–1152. [ PMC free article ] [ PubMed ] [ Google Scholar ]