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Structural characterization of superbug proteins involved in regulating beta-lactam resistance Wilke, Mark Steven 2008

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STRUCTURAL CHARACTERIZATION OF SUPERBUG PROTEINS INVOLVED IN REGULATING β-LACTAM RESISTANCE by MARK STEVEN WILKE B.Sc., University of Guelph, 2002  A Thesis Submitted in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy  in  The Faculty of Graduate Studies (Biochemistry and Molecular Biology)  The University of British Columbia (Vancouver)  April 2008  © Mark Steven Wilke 2008  ABSTRACT β-Lactam antibiotics have been indispensable in treating bacterial infections for more than sixty years and remain the most widely utilized of all antibiotic classes. β-Lactams kill bacteria by interfering with the function of penicillin binding proteins (PBPs), essential enzymes that catalyze the crosslinking of peptidoglycan in the bacterial cell wall. Unfortunately, the widespread use of β-lactams has undermined their effectiveness as chemotherapeutic agents by fueling the evolution and dissemination of multiple resistance mechanisms, including: (1) production of hydrolytic β-lactamase enzymes that inactivate β-lactams, (2) expression of PBPs with low-affinity for β-lactams, and (3) overexpression of multidrug efflux pumps which actively clear β-lactams and other toxic substances from the periplasm of Gram-negative bacteria.  The overall goal of this thesis is the structural  characterization of bacterial proteins involved in regulating β-lactam resistance using X-ray crystallography. Staphylococcus aureus is among the most prevalent and antibiotic-resistant of pathogenic bacteria.  The notorious resistance of S. aureus primarily stems from the  production of β-lactamases and PBP2a, a low-affinity PBP which confers broad-spectrum β-lactam resistance in so-called methicillin-resistant S. aureus (MRSA) strains. Expression of either of these resistance determinants is controlled by a homologous regulatory system utilizing a β-lactam-inducible transmembrane ‘sensor/transducer’ (BlaR1/MecR1) and repressor (BlaI/MecI). This dissertation presents the crystal structure of the BlaR1 sensor domain from S. aureus, determined in its apo form at 1.8 Å resolution and acylated with penicillin G at 2.4 Å. These structures illuminate the active-site features that are responsible for the PBP activity of BlaRS and provide mechanistic insights into the role of BlaRS in ii  detecting β-lactam antibiotics and transducing the binding signal across the bacterial cell membrane. In addition, mutation of a conserved P-X-X-P motif in the L2 loop of BlaR1 was shown to prevent induction of β-lactamase expression in vivo, supporting the hypothesis that the L2 loop plays an important role in signal transduction. The intrinsic resistance of Pseudomonas aeruginosa to a variety of antibiotics including β-lactams is exacerbated in mutant strains that over-express multidrug efflux pumps such as MexAB-OprM. Production of the MexAB-OprM efflux system is negatively regulated by a MarR family repressor, MexR, and several hyper-resistant strains of P. aeruginosa appear to involve mutations in either MexR or additional regulatory factors which influence the activity of MexR. The allosteric effectors of MarR proteins are typically low molecular weight lipophenolic compounds. This dissertation confirms that MexR is uniquely modulated by a 53 amino acid protein called ArmR. Electromobility gel shift assays and isothermal titration calorimetry demonstrate that a direct protein-protein interaction between MexR and ArmR is responsible for neutralizing the DNA-affinity of MexR for its cognate DNA operator. The allosteric conformational change induced by ArmR-binding was assessed by determining the 1.8 Å crystal structure of MexR double mutant Q106L/A110L (MexRLL) in complex with ArmR residues 29-53 (ArmRC). This structure reveals that ArmR induces a dramatic conformational change which repositions the MexR DNA-binding lobes into an orientation that is incompatible with binding DNA.  iii  TABLE OF CONTENTS ABSTRACT................................................................................................... ii TABLE OF CONTENTS ............................................................................ iv LIST OF TABLES ...................................................................................... vii LIST OF FIGURES ................................................................................... viii ABBREVIATIONS ....................................................................................... x ACKNOWLEDGEMENTS ...................................................................... xiii CO-AUTHORSHIP STATEMENT .......................................................... xv CHAPTER 1 – Introduction ........................................................................ 1 1.1 DISCOVERY AND DEVELOPMENT OF β-LACTAM ANTIBIOTICS ................................ 1 1.2 THE ACTION OF β-LACTAMS ON THE BACTERIAL CELL WALL ................................ 4 1.3 PENICILLIN BINDING PROTEINS .................................................................................. 9 1.4 β-LACTAM RESISTANCE: ORIGINS, MECHANISMS AND REGULATION ....................... 14 1.4.1 Origins of Antibiotic Resistance ................................................................................ 14 1.4.2 Mechanisms of β-Lactam Resistance ........................................................................ 15 The β-lactamases ................................................................................................ 15 Resistant PBPs .................................................................................................... 22 Multidrug efflux pumps...................................................................................... 24 Other β-lactam resistance mechanisms............................................................... 27 1.4.3 Regulation of β-Lactam Resistance ........................................................................... 29  1.5 OBJECTIVES OF THESIS ............................................................................................. 32  CHAPTER 2 – The β-Lactam Sensor of BlaR1....................................... 34 2.1 INTRODUCTION ....................................................................................................... 34 2.2 METHODS ................................................................................................................ 38 2.2.1 Cloning, Protein Expression, and Purification............................................................ 38 2.2.2 Crystallization and Data Collection............................................................................ 39  iv  2.2.3 Structure Solution and Refinement............................................................................. 40 2.2.4 Static Light Scattering ................................................................................................ 41 2.2.5 Mass Spectrometry ..................................................................................................... 42  2.3 RESULTS .................................................................................................................. 42 2.3.1 Functional Characterization of Recombinant BlaRS................................................... 42 2.3.2 Overall Fold and Oligomerization State of BlaRS ...................................................... 43 2.3.3 BlaRS Active Site Architecture and the Benzylpenicillin Adduct .............................. 46  2.4 DISCUSSION............................................................................................................. 50 2.4.1 The Role of Lys392 as the General Base in Acylation............................................... 50 2.4.2 The Nζ-Carboxylation of Lys392 .............................................................................. 52 2.4.3 Formation of the Stable Penicilloyl Adduct .............................................................. 54 2.4.4 Structural Differences Between Apo- and Acyl-BlaRS .............................................. 58 2.4.5 Transmembrane Signal Transduction ........................................................................ 61  CHAPTER 3 – The BlaR1 L2 Loop & Signal Transduction ................. 64 3.1 INTRODUCTION ....................................................................................................... 64 3.2 METHODS ................................................................................................................ 65 3.2.1 Expression and Purification of 15N and 2H/15N/13C-Labelled BlaRS........................... 65 3.2.2 NMR Data Collection and Assignment ..................................................................... 66 3.2.3 BlaR1 Mutant Constructs ........................................................................................... 67 3.2.4 β-Lactamase assay ...................................................................................................... 67 3.2.5 Cloning and Purification of the L2 Loop ................................................................... 68 3.2.6 Pull-Down and Crosslinking Assays with GFPL2SOL and BlaRS............................. 73  3.3 RESULTS .................................................................................................................. 74 3.3.1 Pull-Down and Crosslinking Assays with GFPL2SOL and BlaRS............................. 74 3.3.2 Constructs of the L2 Loop and Interaction with BlaRS .............................................. 77 3.4 DISCUSSION ....................................................................................................................... 82 3.4.1 The Role of the L2 Loop in BlaR1 Signal Transduction ............................................ 82  CHAPTER 4 – The Modulation of MexR by ArmR ............................... 84 4.1 INTRODUCTION ....................................................................................................... 84 4.2 METHODS ................................................................................................................ 87 4.2.1 Cloning and Purification of MexR and ArmR............................................................ 87  v  4.2.2 Isothermal Titration Calorimetry (ITC) of MexR-ArmR ........................................... 89 4.2.3 Electromobility Shift Assay (EMSA) of MexR-ArmR .............................................. 90 4.2.4 Limited Trypsinolysis of ArmR and MexR-ArmR..................................................... 90 4.2.5 Expression and Purification of 15N/13C-Labelled ArmR............................................. 91 4.2.6 NMR Data Collection and Assignment ...................................................................... 91 4.2.7 Crystallization and Structure Determination .............................................................. 92  4.3 RESULTS .................................................................................................................. 93 4.3.1 MexR-ArmR Interaction............................................................................................. 93 4.3.2 ArmR Modulates MexR DNA Binding Affinity ........................................................ 93 4.3.3 Isolation of the Minimal Peptide for Binding MexR .................................................. 94 NMR spectral assignments of ArmR .................................................................. 95 Limited trypsinolysis of ArmR in complex with MexR ..................................... 96 ArmR truncations................................................................................................ 98 4.3.4 Crystal Structure of the MexRLL-ArmRC Complex .................................................. 100 Architecture of the MexRLL-ArmRC complex .................................................. 100 Interactions between ArmRC and MexRLL ........................................................ 101  4.4 DISCUSSION ........................................................................................................... 104 4.4.1 Structural Symmetry within ArmR........................................................................... 105 4.4.2 MexR and ArmR Mutants ........................................................................................ 106 4.4.3 ArmR-Induced Conformational Change................................................................... 107 4.4.4 Allosteric Mechanism of MexR Anti-Repression .................................................... 109 4.4.5 Comparison with Other MarR Family Members...................................................... 111  CHAPTER 5 – Conclusions and Future Directions .............................. 115 5.1 THE BLAR1 SENSOR/TRANSDUCER FROM S. AUREUS ......................................... 115 5.2 THE MEXR ANTI-REPRESSOR ARMR FROM P. AERUGINOSA .............................. 118  REFERENCES.......................................................................................... 121 APPENDIX – Publications Arising from Graduate Work................... 149 APPENDIX II – Assigned NMR Peak Lists for Apo-BlaRS ................. 150  vi  LIST OF TABLES Table 1.1: Major subclasses of β-lactam antibiotics ....................................................................... 3 Table 2.1: Crystallographic data and refinement statistics ........................................................... 44 Table 3.1: Description of various L2 loop constructs.................................................................... 69 Table 3.2: Cloning of the L2 loop.................................................................................................. 70 Table 3.3: Summary of mutations and their effects on CBAP inducible β-lactamase expression assayed by hydrolysis of 100 µM cephaloridine ..................................... 77 Table 3.4: Expression of various L2 loop constructs and when applicable the testing of an interaction with BlaRS ........................................................................................ 79 Table 4.1: Data collection and refinement statistics. .................................................................. 101  vii  LIST OF FIGURES Figure 1.1: Chemical structure of 6-amino-penicillanic acid (6-APA) ............................................. 2 Figure 1.2: Crosslinking of peptidoglycan in the bacterial cell wall ............................................... 6 Figure 1.3: Structure and mechanism of PBPs ........................................................................... 11 Figure 1.4: β-Lactam antibiotics as suicide substrates for the PBPs .......................................... 13 Figure 1.5: Graphical summary of β-lactam resistance mechanisms and their regulation .......... 16 Figure 1.6: Representative serine β-lactamase CTX-M9 (class A) .............................................. 18 Figure 1.7: Representative class B β-lactamase CphA ................................................................ 21 Figure 2.1: Regulation systems controlling the expression of β-lactamase and PBP2a (shown in large brackets) .......................................................................................... 37 Figure 2.2: Structure of apo-BlaRS ............................................................................................... 46 Figure 2.3: Amino acid sequence alignment of BlaRS with the BlaR1 sensor domains of Bacillus licheniformis (BlaR1-Bl), Staphylococcus haemolyticus (BlaR1-Sh), the MecR1 sensor domains of S. aureus (MecR1-Sa), Staphylococcus epidermis (MecR1-Se), and a representative class D β-lactamase OXA-10 from Pseudomonas aeruginosa (OXA10-Pa) ................................................................................................................ 48 Figure 2.4: Active site of (a) apo- and (b) acyl-BlaRS ................................................................... 49 Figure 2.5: Electron density of Lys392 and PenG adduct ........................................................... 49 Figure 2.6: Comparison of the apo-BlaRS active site with representative β-lactamases and a PBP ........................................................................................................... 51 Figure 2.7: Acylation schemes considering Lys392 as (a) non-carboxylated and (b) carboxylated .......................................................................................................... 55 Figure 2.8: Acylation schemes considering protonation of Lys392 at (a) Nζ facilitating decarboxylation or (b) Oθ facilitating deacylation (modified from Cha and Mobashery, 2007)....................................................................................................... 59 Figure 2.9: Overlay of apo- and acyl-BlaRS .................................................................................. 61 Figure 3.1: Overlapped 1H/15N-TROSY-HSQC spectra showing the differences in chemical shifts between apo-BlaRS (red) and BlaRS acylated by PenG (blue) ........................ 75  viii  Figure 3.2: The crystal structure of PenG-acylated BlaRS demonstrating the proximity of the PenG adduct to residues corresponding to successfully assigned 1  H/15N-HSQC peaks ................................................................................................... 76  Figure 3.3: GFPL2SOL fluorescence and pull down assay with BlaRS........................................ 80 Figure 3.4: GFPL2SOL cross-linking assays with BlaRS.............................................................. 81 Figure 4.1: Schematic detailing the regulation MexAB-OprM-mediated efflux ............................ 86 Figure 4.2: Binding of ArmR to MexR as measured by ITC ......................................................... 94 Figure 4.3: MexR-ArmR complex formation abrogates MexR-DNA binding ................................ 95 Figure 4.4: Assigned 1H/15N-HSQC spectrum of the 15N/13C-ArmR fragment, residues 25-53........................................................................................................................... 97 Figure 4.5: Secondary structure propensity (SSP) of free 15N/13C-ArmR25-53............................... 98 Figure 4.6: MALDI-TOF mass spectra displaying limited tryptic digests of (a) ArmR with BSA and (b) ArmR with MexR .................................................................................... 99 Figure 4.7: Binding affinity of MexR with N- and C-terminally truncated ArmR constructs ........ 100 Figure 4.8: The crystal structure of MexRLL in complex with ArmRC .......................................... 102 Figure 4.9: Interactions between MexRLL and ArmRC ................................................................ 103 Figure 4.10: Overlapping binding sites reveal structural symmetry in the ArmR C-terminus..... 104 Figure 4.11: The binding of ArmRC induces a conformational change in MexRLL...................... 108 Figure 4.12: Mechanism of anti-repression ................................................................................ 111  ix  ABBREVIATIONS ΔH  binding enthalpy change  ΔS  binding entropy change  3D  three-dimensional  6-APA  6-aminopenicillanic acid  7-APA  7-aminocephalosporanic acid  A280  absorbance at 280 nm  ArmRC  ArmR-derived peptide spanning residues 29-53  BisTris  bis(2-hydroxyethyl)-imino-tris-(hydroxymethyl)-methane  BlaRP  BlaR1 protease domain  S  BlaR  BlaR1 sensor domain  BSA  bovine serum albumin  BsOhrR  Bacillus subtilus OhrR  CBAP  2-(2′-carboxyphenyl)-benzoyl-β-aminopenicillanic acid  CBD  chitin-binding domain  CNS  Crystallography and NMR System  Da  dalton  DAP  LL-diaminopimelic acid  DMSO  dimethyl sulfoxide  DNA  deoxyribonucleic acid  DNAop  double-stranded operator DNA  DTT  dithiothreitol  DSS  disuccinimidyl suberate  EDTA  ethylene-diamine tetra-acetic acid  ELISA  enzyme-linked immunosorbent assay  EMSA  electromobility shift assay  ESBL  extended spectrum β-lactamase  FT-IR  Fourier-transformed infrared  GlcNAc  N-acetylglucosamine  HEPES  4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid  HPLC  high-performance liquid chromatography x  HSQC  heteronuclear single quantum coherence  HMM  high molecular mass  IM  inner membrane  IPTG  isopropyl-β-D-thiogalactopyranoside  ITC  isothermal titration calorimetry  Kd  dissociation equilibrium constant  kDa  kilodalton  LMM  low molecular mass  LB  Luria Bertani medium  MAD  multiwavelength anomalous diffraction  MALDI-TOF  matrix-assisted laser desorption ionization time of flight  MexRLL  MexR Q106A/A110L mutant  MPD  2-methyl-2,4-pentanediol  MRSA  methicillin-resistant Staphylococcus aureus  MS  mass spectrometry  MurNAc  N-acetylmuramic acid  N  binding stoichiometry  NMR  nuclear magnetic resonance  NSLS  National Synchrotron Light Source  OD600  optical density at 600 nm  OM  outer membrane  PAGE  polyacrylamide gel electrophoresis  PBP  penicillin-binding protein  PCR  polymerase chain reaction  PDB  Protein Databank  PEG  polyethylene glycol  PenG  penicillin G (i.e. benzylpenicillin)  PG  peptidoglycan  PRSP  penicillin-resistant Streptococcus pneumoniae  rms  root mean squared  rmsd  root mean square deviation xi  RND  Resistance-Nodulation-Division  SDS  sodium dodecylsulfate  SeMet  selenomethionine  ssDNA  single-stranded DNA  ssp.  species (i.e. within a particular genus)  SSP  secondary structure propensity  TCEP  tris-(carboxyethyl)phosphine hydrochloride  Tris  2-amino-2-hydroxymethyl-1,3-propanediol  TROSY  transverse relaxation optimized spectroscopy  UV  ultraviolet  v/v  unit volume (mL) per unit volume (mL)  w/v  unit weight (g) per unit volume (mL)  XcOhrR  Xanthomonas campestris OhrR  xii  ACKNOWLEDGEMENTS First of all, I would like to thank my parents for their unconditional support during my undergraduate and graduate degrees. In addition, I am indebted to Candice Mathany for her exuberant encouragement and support while writing this thesis. Candice is a constant inspiration to me and her love and attention sustained me during every page of this manuscript. This thesis would not have been possible without the support of my supervisor, Dr. Natalie Strynadka. Natalie provided me with a tremendous learning environment and her guidance in my research has been invaluable. I would also like to thank my supervisory committee members, Drs. Grant Mauk and Lawrence McIntosh, who provided me with valuable insights and direction for my work. This work is greatly enhanced by excellent collaborations with Dr. Henry Chambers in the Department of Medicine at the University of California, San Francisco and Dr. Keith Poole in the Department of Microbiology and Immunology at Queen’s University. In addition, I would like to thank Dr. Markus Heller from the Dr. Lawrence McIntosh laboratory for his assistance in NMR collection and analysis as well as Dr. Louise Creagh for her repeated assistance in performing isothermal titration calorimetry experiments. I would like to extend my appreciation for the assistance I obtained from all members of the Strynadka laboratory, both past and present. Special thanks to Liza deCastro, Tanya Hills and Richard Pfuetzner for their technical advice and laboratory training in cloning and protein purification. I would like to thank Drs. Andrew Lovering and Daniel Lim for teaching me protein crystallography and my benchmate Michael Gretes for keeping me sane on project BlaR with heated political discussions.  xiii  I am extremely grateful for the financial support from the Natural Sciences and Engineering Research Council of Canada, the Canadian Institutes of Health Research, and the Michael Smith Foundation for Health Research.  I would also like to extend my  appreciation to Drs. Natalie Strynadka, Lawrence McIntosh, Andrew Lovering, Lynne Howell, and Tim Zacharewski for their time and effort over the years providing me with reference letters.  xiv  CO-AUTHORSHIP STATEMENT Chapters 1 to 4 of this thesis describe work that was previously published. Manuscripts for these chapters were written by myself and revised by my supervisor Dr. Natalie Strynadka. Chapter 1 contains portions from a manuscript published in Current Opinion in Microbiology [Wilke, M. S., Lovering, A. L., Strynadka, N. C. (2005) β-Lactam antibiotic resistance: a current structural perspective. Curr Opin Microbiol 8(5), 525-33]. In the original publication, the text and figures for the β-lactamase sections were prepared by me, the text and figures for the PBP section were prepared by Dr. Andrew Lovering, the text for the introduction and efflux pump sections was prepared by Dr. Natalie Strynadka, and the figure for the introduction was prepared jointly by myself and Dr. Andrew Lovering. Chapters 2 and 3 contain portions from a manuscript published in the Journal of Biological Chemistry [Wilke, M. S., Hills, T. L., Zhang, H.-Z., Chambers, H. F. and Strynadka, N. C. J. (2004) Crystal structures of the apo and penicillin-acylated forms of the BlaR1 β-lactam sensor of Staphylococcus aureus. J Biol Chem 279(45), 47278-87]. Dr. Henry Chambers provided the initial full-length construct of BlaR1. Tanya Hills designed the sensor domain construct and developed an initial protein purification protocol.  I  modified the purification protocol and performed the crystallization, data collection, structure determination, and refinement. In the original publication, all text and figures were prepared by me except for the sections pertaining to the in vivo mutagenesis of BlaR1 which was performed by Drs. Hong-Zhong Zhang and Henry Chambers. Chapter 4 contains portions from a manuscript published in the Journal of Bacteriology [Daigle, D. M., Cao, L., Fraud, S., Wilke, M. S., Pacey, A., Klinoski, R., Strynadka, N. C., Dean, C. R., and Poole, K. (2007). A protein modulator of multidrug xv  efflux gene expression in Pseudomonas aeruginosa. J Bacteriol 189(15), 5441-51]. Twohybrid screens were prepared by Lily Cao and Sebastian Fraud.  Isothermal titration  calorimetry was performed by Denis Daigle (although in Chapter 4, isothermal titration calorimetry was performed by Dr. Louise Creagh). The electromobility shift assay was prepared by me. In the original publication, all text and figures were prepared by Denis Daigle, Lily Cao and Keith Poole except for the figures pertaining to the electromobility shift assay and structure of MexR. The majority of the work presented in Chapter 4 (e.g. limited proteolysis, NMR, crystallization and X-ray crystallography) has not been previously published and is the source of a future research article.  xvi  CHAPTER 1 – Introduction  1.1  DISCOVERY AND DEVELOPMENT OF β-LACTAM ANTIBIOTICS For billions of years, microbes have been developing chemical and biological  weapons to gain advantage within their respective ecological niches. With Sir Alexander Fleming’s discovery of penicillin in Penicillium mould (Fleming, 1929), humankind began to engage microbial pathogens in a biochemical arms race which bears an increasingly uncertain outcome due to the rapid and widespread evolution of antibiotic resistance. Penicillins—along with cephalosporins, carbapenems, and monobactams—constitute a class of antibiotics called β-lactams (Table 1.1) and are considered the most widely used antimicrobial drugs by virtue of their unmatched efficacy and safety profiles (Essack, 2001). The common feature of β-lactam antibiotics is the β-lactam ring, an amide-containing fourmembered ring with bactericidal activity (Crowfoot et al., 1949). The efficacy of penicillin as a systemic chemotherapeutic agent for bacterial infections was first demonstrated in mice by Chain, Florey and colleagues in 1940 (Chain et al., 1993) and then in humans in 1941 (Abraham et al., 1992). These successes motivated innovations in fermentation for large scale production of penicillin and modification of the acyl side-chain by the addition of precursor compounds to the fermentation medium (Rolinson, 1998). However, the toxicity of side-chain precursors limited the diversity of penicillin derivatives until the ‘penicillin nucleus’ (6-APA or 6-aminopenicillanic acid) was isolated by fermenting Penicillium in the absence of precursor compounds (Figure 1.1). 6-APA was later produced in higher yields for the construction of penicillin derivatives by removing the penicillin side-chain by enzymatic deacylation (Rolinson et al., 1960). The 1  availability of 6-APA provided the opportunity to develop semisynthetic penicillins with improved oral absorption, broader spectrums of activity, and enhanced stability against penicillin-resistant bacteria.  Similarly, the subsequent discovery of cephalosporin C in  Cephalosporium acremonium by Giuseppe Brotzu (Brotzu, 1948) and the characterization of its structure (Abraham and Newton, 1961; Hodgkin and Maslen, 1961) led to the isolation of the cephalosporin nucleus (7-ACA or 7-aminocephalosporanic acid) which enabled construction of semisynthetic cephalosporin derivatives (Loder et al., 1961).  The  carbapenems, derived from Streptomyces cattleya (Brown et al., 1976; Weaver et al., 1979), the monobactams, derived from Chromobacterium violaceum (Sykes and Bonner, 1985), and the β-lactamase-inhibiting clavams (Brown et al., 1976), derived from Streptomyces clavuligerus, further expanded the power of the β-lactams as antimicrobial drugs .  1  H2N  6 7  O  S  5  N  2 3  4  O  O  -  Figure 1.1: Chemical structure of 6-amino-penicillanic acid (6-APA). The absence of an acyl side chain in 6-APA (versus penicillin G in Table 1.1) permits modification of the free amino group and construction of a large variety of semisynthetic penicillins with altered properties.  2  Table 1.1: Major subclasses of β-lactam antibiotics e.g. Penicillin G  Penicillins H N  R  H N  S  O O NH  O  S  N  e.g. Cefipime (4th Generation) H2N  O  Other Cephalosporins  COOH  H N  R1  dicloxacillin, nafcillin,  COOH  ampicillin, amoxicillin,  Cephalosporins  O  oxacillin, cloxacillin,  N  COOH  O  penicillin V, methicillin,  S  O  N  R  Other Penicillins  1st Generation: cepalothin,  N  S S  R2  N O COOH  N  N O  O  cephapirin, cefazolin,  S  H N  +  N  cephradine, cephaloridine  C  2nd Generation: cefamandole,  COOH  O  cefuroxime, cephalexin,  Carbapenems R1  e.g. Imipenem OH  N  COOH  HN COOH  O  Monobactams HOOC  O  O  N  H2N  O S  β-Lactamase Inhibitors  none commercially available  N O  OH  O  R2  N O  COOH  3  SO 3H  Other β-Lactamase  e.g. Clavulanate  O  R1  Other Monobactams  N SO3H  N  NH  H N  N O  thienamycin  e.g. Aztreonam  H N  R  meropenem  S  R2  N O  O  Other Carbapenems  Inhibitors  1.2  THE ACTION OF β-LACTAMS ON THE BACTERIAL CELL WALL The cytoplasmic membrane of a bacterial cell is encased in a mesh-like exoskeleton  called the sacculus (or cell wall), which endows bacteria with mechanical strength and maintains cell shape. Without this cell wall, a bacterium would be unable to withstand the osmotic stress across its cytoplasmic membrane, lyse and die. It has long been recognized that β-lactam antibiotics target biosynthesis of the sacculus (Tipper and Strominger, 1965), but until recently it was not clear exactly how these drugs induce bacterial death. As a bacterium grows and divides, its cell wall must adapt accordingly. A large multienzyme complex containing both synthases and hydrolases is thought to coordinate this delicate process of remodeling the sacculus so as to prevent bacteriolysis (Holtje, 1996). Early research demonstrated that sub-lethal doses of penicillin cause abnormal cell morphology and incomplete fission, whereas lethal doses result in cell lysis (Gardner, 1940). Thus, it was widely believed that by inhibiting cell wall biosynthesis, β-lactams induce a fatal attack of cell wall hydrolases (i.e. bacterial lytic enzymes), which dismantle the sacculus to cause lysis and death. However, electron microscopic studies documenting penicillin-induced death in S. aureus have shown that bacteriolysis represents only a postmortem process (Giesbrecht et al., 1998).  Instead, the killing effect of penicillin is apparently a result of induced  morphogenetic defects of the sacculus at the cell division plane that become fatal during cell separation. In normal bacterial reproduction, a cell wall septum forms to close off the two daughter cells prior to punching pores in the sacculus along the cell division plane in the initial stages of cell separation.  However, when these pores develop on top of a  defective/incomplete septum (due to penicillin-mediated impairment of cell wall synthesis), the high internal pressure of the protoplast abruptly kills the bacterium via ejection of a 4  portion of cytoplasm during attempted cell separation. The disintegration of the bacterial cell via postmortem bacteriolysis then results from the shrinkage of the dead cell and the perturbation of its cytoplasmic membrane. The sacculus is composed of crosslinked strands of peptidoglycan, a biopolymer consisting  of  alternating  residues  of  N-acetylmuramic  acid  (MurNAc)  and  N-acetylglucosamine (GlcNAc) linked by β-(1,4)-glucosidic bonds and modified with a short peptide stem of four or five amino acids (Walsh, 2003). The peptide stem consists of L-Ala-γ-D-Glu-L-Lys/DAP-D-Ala(-D-Ala) and is attached to the lactyl group of MurNAc  residues (Figure 1.2). In Gram-negative and some Gram-positive bacteria, the third amino acid in this peptide is LL-diaminopimelic acid (DAP), whereas Gram-positive species like S. aureus use L-Lys preceded by an α-amidated D-Glu residue. Peptidoglycan strands are crosslinked between the free amino group of L-Lys/DAP and the penultimate D-Ala of a separate strand; additional residues can be involved in the crosslink as an interbridge (e.g. pentaglycine in S. aureus). Extensive crosslinking creates a massive peptidoglycan net (also called murein) which encompasses the entire bacterial cell to form the sacculus. The murein sacculus of Gram-positive bacteria is much thicker than in Gram-negative species and constitutes a surface organelle for the display of carbohydrates and proteins by analogy with the outer membrane of Gram-negative bacteria (Lee and Schneewind, 2001). The 3D architecture of peptidoglycan is presently unknown. This is principally because the sacculus of a living cell is in a state of perpetual biosynthesis (i.e. assembly, disassembly and turnover), which produces heterogeneous samples that are unconducive to structural analysis. Several models have been deduced from indirect evidence and largely differ with respect to the orientation of the glycan strands relative to the cytoplasmic  5  GlcNAc MurNAc  OH  HO  OH  O  O NH O  GlcNAc  OH  O  O  O  HO  NH  O  NH  O  NH  L-Ala  O  O  O  O O  HN COOH  D-Glu  OH  O  DAP  HO  O H2N  NH O  COOH O  OH  O  O  HN  D-Ala-D-Ala  OH  O  O  NH  NH  O  NH  COOH  O  HO  NH  O  NH  O  O  O  O O  HN COOH O NH  D-Ala  O  H N  HN COOH O  O COOH H2N  NH  HN O  COOH  NH O COOH HN O NH O  O O  O  O NH  NH HO  O  O O  O  NH  HO  O  O OH  O  OH  OH  Figure 1.2: Crosslinking of peptidoglycan in the bacterial cell wall. Abbreviations: GlcNAc, N-acetylglucosamine; MurNAc, N-acetylmuramic acid; DAP, LL-diaminopimelic acid.  6  membrane. The “classical” model proposes that glycan strands run in an extended linear conformation parallel to the cytoplasmic membrane and perpendicular to the long axis of rod-shaped cells (Holtje, 1998). A variation of this model contends that in the stress-bearing wall, successive MurNAc-GlcNAc disaccharides rotate 90° and thus crosslink in a hexagonal “chicken-wire” network (Koch, 1998). Either of these models results in the formation of two-dimensional peptidoglycan layers from the crosslinking of glycan strands.  Thus a  monolayer of peptidoglycan produces the relatively thin cell wall of Gram-negative bacteria whereas the thicker Gram-positive sacculus consists of multiple layers crosslinked in threedimensions. The “scaffold” model uses a perpendicular orientation of glycan strands relative to the cytoplasmic membrane (Dmitriev et al., 2005). The original form of this model assumes that glycan strands adopt a helical conformation that presents four peptide side chains per turn of the helix (as in the chicken-wire model). However, the recent NMR structure of a synthetic fragment of peptidoglycan revealed a helical conformation of three MurNAc-GlcNAc repeats per turn (Meroueh et al., 2006). This suggests a glycan strand with a maximum of three peptide crosslinks per helical turn, which would produce a compelling honeycomb structure when oriented orthogonal to the cytoplasmic membrane. Peptidoglycan biosynthesis can be separated into two stages: (1) construction of disaccharide-peptide monomer units (i.e. Lipid II) in the cytoplasm and (2) polymerization and crosslinking of these monomer units on the outside surface of the cytoplasmic membrane (van Heijenoort, 1998; van Heijenoort, 2001). Assembly of Lipid II begins with conversion of fructose-6-phosphate to uridine-5’-pyrophosphate-GlcNAc (UDP-GlcNAc) in four successive cytoplasmic steps catalyzed by GlmS, GlmM and bifunctional GlmU. MurA and MurB form uridine-5’-pyrophosphate-MurNAc (UDP-MurNAc) from UDP-GlcNAc, at  7  which point amino acid residues are then added by the action of four enzymes, MurC to MurF.  MurY next transfers the phospho-MurNAc-pentapeptide moiety from the UDP-  MurNAc-peptide precursor to the membrane-bound acceptor undecaprenyl phosphate to yield Lipid I. The synthesis of Lipid II is completed by the MurG-mediated addition of GlcNAc to Lipid I. Depending on the bacterial species, the pentapeptide substituent may be modified during the formation of these lipid intermediates (e.g. amidation or the addition of interbridge amino acids). Lipid II is then translocated across the cytoplasmic membrane by an unknown mechanism where glycosyltransferases catalyze polymerization of the glycan strands by transfer of the disaccharide-peptide monomer to the growing peptidoglycan chain. The peptide crosslinks are formed by transpeptidase enzymes which break the peptide bond of a C-terminal D-Ala-D-Ala donor and form a new peptide bond with the amino group of an amino acid acceptor on a separate strand, either L-Lys, DAP, or an interbridge residue. The degree of crosslinking varies between bacteria [approximately 40% in E. coli (Glauner et al., 1988)] and is continually modified by the activity of murein synthases and hydrolases as required for growth and cell division. Transpeptidation is the final step in the biosynthesis of peptidoglycan and is absolutely essential to the structural integrity of the sacculus and the survival of the bacterial cell. The enzymes that carry out this critical step are the targets of β-lactam antibiotics, which kill bacteria by inhibiting the formation of crosslinks between peptidogylcan strands.  8  1.3  PENICILLIN-BINDING PROTEINS Peptidoglycan crosslinking is catalyzed by transpeptidase enzymes called penicillin-  binding proteins (PBPs), so named due to their ability to form stable covalent complexes with β-lactams. Bacteria have multiple PBPs which play various roles in modulating the sacculus (Macheboeuf et al., 2006; Massova and Mobashery, 1998). Not all PBPs are involved in peptidoglycan synthesis and a particular PBP may exhibit DD-transpeptidase, DD-carboxypeptidase, DD-endopeptidase, or glycosyltransferase activities.  PBPs are  typically divided into two groups based on amino acid sequence: low molecular mass (LMM; 20-50 kDa) or high molecular mass (HMM; 60-120 kDa).  The soluble or membrane-  associated LMM PBPs are classified according to their specific activities (classes A – C). Of these, the class B enzymes appear to be involved in peptidoglycan turnover and cell shape by controlling the degree of crosslinking in the cell wall. Among the LMM PBPs, the class A enzymes are DD-transpeptidases, the class C enzymes are DD-endopeptidases, and the class B enzymes may exhibit both DD-transpeptidase and DD-carboxypeptidase activities. Synthesis of new peptidoglycan (as in cell division) is performed by the membrane-anchored HMM PBPs, which can be sub-divided into bifunctional glycosyltransferase/transpeptidase enzymes (class A) and monofunctional transpeptidase enzymes (class B). Class C HMM PBPs are transmembrane receptors involved in sensing β-lactams and signaling the production of proteins involved in antibiotic resistance (Hardt et al., 1997; Zhang et al., 2001). Structural studies have provided valuable insights into the catalytic mechanisms of PBPs and their inhibition by β-lactams. Representative crystal structures from class A (McDonough et al., 2002; Rhazi et al., 2003; Sauvage et al., 2005) and class B LMM PBPs 9  (Morlot et al., 2005; Nicola et al., 2005) as well as class A (Contreras-Martel et al., 2006; Lovering et al., 2007; Macheboeuf et al., 2005; Yuan et al., 2007) and class B HMM PBPs (Gordon et al., 2000; Lim and Strynadka, 2002; Sauvage et al., 2002) have shown each of these enzymes to possess a transpeptidase domain with a similar fold and active site architecture containing three highly conserved motifs: S-X-X-K, S/Y-X-N, and H/K-T/S-G (Fisher et al., 2005). The catalytic mechanism (Figure 1.3) for this domain is proposed to begin with activation of a serine nucleophile by a general base lysine (both residues found within the S-X-X-K motif) and attack of the peptide carbonyl by the nucleophilic serine to form an acyl-enzyme intermediate (Rhazi et al., 2003). The amino group of the peptide acceptor is then activated for nucleophilic attack on the acyl-serine ester by the serine/tyrosine of the S/Y-X-N motif (Silvaggi et al., 2003), resulting in deacylation and acyl transfer to the peptide acceptor. In carboxypeptidase reactions, water substitutes for the peptide acceptor in deacylation, preventing future crosslinking by hydrolysis of the peptide stem. The glycosyltransferase activity of the class A HMM PBPs is contained on a distinct domain positioned after the transmembrane anchor and separated from the C-terminal transpeptidase domain by a flexible linker (Figure 1.3). The polymerization mechanism is proposed to use two catalytic glutamate residues. The first glutamate deprotonates the 4-OH group of the Lipid II acceptor, which subsequently attacks C1 of the growing glycan-chain donor. An additional glutamate may facilitate this process by direct protonation of the phospho-sugar bond or metal-mediated stabilization of the pyrophosphate group of the glycan-chain (Lovering et al., 2007). The class B HMM PBPs also contain a non-penicillinbinding domain positioned between the transmembrane anchor and transpeptidase domain. This domain shares no structural resemblance to the glycosyltransferase domain of class A  10  S398  K401  D-Ala  D-Ala  Y454  R HN H  acceptor peptide–NH2  H N  H O  R  O  -  O  HN  PBP(Ser)–OH  O  H  PBP  O  +  R  H  HN  NH2 O  O  (a)  H N  -  (b)  acceptor peptide  +  H O  PBP(Ser)–OH  E114  Lipid II  growing glycan-chain E171  (c) Figure 1.3: Structure and mechanism of PBPs. (a) Structure of bifunctional PBP2 with catalytic residues shown as red spheres (PDB code 2OLU). (b) Transpeptidation—Mechanism and close up view of active site (PDB code 1IKG, R61 DD-peptidase from Streptomyces R61) bound to peptidoglycan substrate fragment containing D-Ala-D-Ala. Active site residues (PBP2 numbering) and substrate are colored by atom type (protein C atoms, green; substrate C, magenta; O, red; and N, blue). (c) Transglycoslation—Mechanism and close up view of active site (PBP2 from S. aureus) modeled with growing glycan-chain donor (left side) and the Lipid II acceptor (right side) [reproduced from (Lovering et al., 2007)]. Active site residues and substrates are colored by atom type (protein C atoms, blue; substrate C, yellow; O, red; N, dark blue; and P, orange).  11  HMM PBPs, but has been suggested to participate in protein-protein interactions and targeting of the class B HMM PBPs to the divisome, a putative complex of cytoplasmic, membrane, and periplasmic proteins involved in septum formation during cell division (Marrec-Fairley et al., 2000). Alternatively, the non-penicillin-binding domain may simply serve as a pedestal to position the transpeptidase domain at a comparable distance from the cytoplasmic membrane as their bifunctional HMM counterparts (Macheboeuf et al., 2006). The power of β-lactam antibiotics derives from their structural similarity to the terminal D-Ala-D-Ala of peptidoglycan stem peptides (Figure 1.4) (Tipper and Strominger, 1965). The highly reactive amide bond of the β-lactam ring substitutes for the peptide bond between D-Ala-D-Ala and the carboxylate group at the C3 position of penicillin is equivalent to the C-terminal carboxylate of the peptide stem. According to the catalytic mechanism for transpeptidation, acylation is accompanied by liberation of D-Ala as the leaving group. While acylation with penicillin cleaves the amide bond of the β-lactam ring, the leaving group remains covalently attached and sterically blocks nucleophilic attack on the penicilloyl-serine ester by either a peptide acceptor or water (Ghuysen et al., 1986). This effectively traps the PBP as an acyl-enzyme intermediate as recyclization of the strained four-membered β-lactam ring is highly unfavoured.  12  (a)  (b)  S454 N O  N H  S398 O  O  K401  PBP PBP(Ser)  PBP(Ser)—OH  stable acyl-PBP  (c) Figure 1.4: β-Lactam antibiotics as suicide substrates for the PBPs. 3D models of (a) the natural substrate of the transpeptidase active site, D-Ala-D-Ala and (b) the suicide substrate, penicillin G. Carbon atoms in penicillin G mimicking D-Ala-D-Ala are coloured yellow (other C atoms, white; N, blue; O, red; and S, green). (c) Mechanism and close up view of PBP active site (PDB code 1QMF, PBP2x from Streptococcus pneumoniae) covalently-bound to the β-lactam cefuroxime. Active site residues and cefuroxime are colored by atom type (protein C atoms, green; cefuroxime C, magenta; O, red; and N, blue). For consistency, amino acid numbering is from S. aureus PBP2.  13  1.4  β-LACTAM RESISTANCE: ORIGINS, MECHANISMS & REGULATION  1.4.1  Origins of Antibiotic Resistance Despite the tremendous success of antibiotics in controlling bacterial infections,  antibiotic resistance has been increasing at an alarming rate (Levy, 2001). The majority of antibiotics in human use are natural products, developed by microbes as chemical weapons to exterminate neighbouring microbial species.  Auto-protection strategies (i.e. resistance  mechanisms) for these antibiotic-producing bacteria (or fungi) are thought to have coevolved with antibiotic biosynthetic pathways to prevent self-destruction. Antibiotic resistance also evolves in response to environmental exposure. For example, hospitals provide an intensive and essentially constant selective pressure for collecting mutations that cultivate antibiotic resistance. While mutation is the inevitable consequence of replicating a large genome with imperfect DNA polymerases, it is becoming clear that certain antibiotics (including β-lactams) can evoke an SOS response in bacteria (Maiques et al., 2006) which enhances errors during genome replication to facilitate ‘programmed evolution’ of resistance (Cirz et al., 2005). Frighteningly, the SOS response appears to also encourage horizontal gene transfer (Kelley, 2006; Maiques et al., 2006), facilitating rapid exchange of antibiotic resistance genes carried on mobile genetic elements (such as plasmids or transposons) between microbes.  While the evolution of antibiotic resistance is a natural ecological  phenomenon, the emerging crisis is a direct result of misuse and overuse of antibiotics in medicine, agriculture, and other human industries (Levy, 2001; Palumbi, 2001). Accordingly, averting the catastrophic failure of healthcare systems around the world will not only require new innovations in antimicrobial therapies, but also enhanced regulations, surveillance, and awareness (Avorn et al., 2001). 14  1.4.2  Mechanisms of β-Lactam Resistance The β-lactams are a highly effective and extremely important class of antibiotics as  they kill a broad spectrum of bacteria, are relatively inexpensive to produce, and generally well tolerated by patients. In addition, the accessibility of the PBPs as β-lactam targets and the lack of PBP homologs in plants and animals have contributed greatly to the success of the β-lactams as chemotherapeutic agents; β-lactams constitute half of the global antibiotics market (Hall and Barlow, 2004). Unfortunately, the widespread use of β-lactam antibiotics has resulted in the selection and dissemination of resistant bacterial strains. There are three major mechanisms of β-lactam resistance: (1) expression of hydrolytic β-lactamase enzymes that inactivate β-lactams, (2) production of PBPs with reduced affinity for β-lactams, and (3) active efflux of β-lactams from the periplasm of Gram-negative bacteria (Figure 1.5). The β-lactamases. β-Lactamases confer considerable antibiotic resistance to their bacterial hosts by hydrolyzing the amide bond of the four-membered β-lactam ring. The spread of β-lactamase genes has been greatly exacerbated by their integration within mobile genetic elements, including plasmids, transposons, and multidrug resistance cassettes, which bestow resistance genes for a variety of antibiotic classes such as β-lactams, aminoglycosides, macrolides, sulphonamides and chloramphenicol (Weldhagen, 2004). Once expressed, β-lactamases are secreted into the periplasmic space (in Gram-negative bacteria), bound to the cytoplasmic membrane, or excreted (in Gram-positive bacteria). The more than 520 β-lactamases known to date ( are typically organized into four classes (A to D) on the basis of sequence identity (Ambler, 1980). Within each class, β-lactamases are further divided into families (denoted by numerals such  15  Figure 1.5:  Graphical summary of β-lactam resistance mechanisms and their regulation.  Genes (shown as solid arrows) and their respective proteins are matched by colour. CTXM9 and CphA are shown as representatives of the ~50 available serine and metallo-β-lactamase structures. Regulation of PBP2a expression is homologous to BlaR/BlaI.  Abbreviations: gm +/–, Gram-  positive/negative; IM, inner membrane; OM, outer membrane; PG, peptidoglycan.  16  as OXA-1, OXA-2, etc.) whose members differ by only a few amino acid substitutions. Crystal structures of representative β-lactamases from each of these classes have been reported. The serine β-lactamases (i.e. classes A, C and D) share a similar fold and mechanism (Figure 1.6), which involves creation of a serine nucleophile by deprotonation of an active site serine with a general base, nucleophilic attack of the β-lactam ring to form an acylenzyme intermediate (i.e. acylation), and hydrolysis of the intermediate using a general base activated water molecule (i.e. deacylation).  Despite striking structural and mechanistic  similarities, the three classes of serine β-lactamase differ in the type of general base residues employed for acylation and deacylation. The mechanism of class A β-lactamases remains the most controversial despite the availability of multiple high-resolution crystal structures for each step of the reaction coordinate, elegant mechanistic and mutagenesis studies, as well as theoretical simulations. The controversy centers around the identity of the general base which deprotonates Ser70 for acylation. Both Lys73 (Golemi-Kotra et al., 2004; Meroueh et al., 2005; Strynadka et al., 1992; Swaren et al., 1995) and Glu166 via a catalytic water (Chen et al., 2007; Lamotte-Brasseur et al., 1991; Minasov et al., 2002; Nukaga et al., 2003) have been mooted as general base candidates. Deacylation in class A β-lactamases is widely accepted to utilize Glu166 as a general base to activate the catalytic water for hydrolysis of the acyl-enzyme intermediate. Class C β-lactamases are thought to employ Tyr150 as a general base in both acylation and deacylation (Dubus et al., 1996; Lobkovsky et al., 1994; Oefner et al., 1990).  The class D β-lactamases appear to employ the carbamate of  Nζ-carboxylated Lys73 as the general base for acylation and deacylation, a rare post-translational modification observed previously in the structures of rubisco 17  K73  S70  E166  (b)  (a) Scheme 1 – Lys73 as general base  Scheme 2 – Glu166 as general base  (c) Figure 1.6:  Representative serine β-lactamase CTX-M9 (class A). (a) Cartoon of the CTX-M9  fold, shown acylated with cefoxitin (PDB code 1YMX). (b) Close-up view of the CTX-M9 active site with several active site residues indicated as sticks, colored by atom type (C atoms, yellow; O, red; N, blue; and S, orange).  The covalently-bound cefoxitin is shown with purple carbon atoms to  distinguish it from the protein carbons displayed in yellow. An active site water molecule is indicated with a cyan sphere. (c) Proposed mechanisms for the enzymatic hydrolysis of a β-lactam [catalytic scheme reproduced from (Nukaga et al., 2003)].  18  (Cleland et al., 1998; Lorimer and Miziorko, 1980), urease (Jabri et al., 1995), and phosphodiesterase (Benning et al., 1995). The reaction scheme of the serine β-lactamases for β-lactam hydrolysis closely resembles that of the PBPs with the exception of an enhanced rate of hydrolysis. In fact, it has been postulated that β-lactamases may have evolved from PBPs (Tipper and Strominger, 1965) and indeed the structural similarities between the PBPs and the class A, C, and D β-lactamases are difficult to ignore (Hall and Barlow, 2004). This conservation of structure exists despite a dearth of sequence identity beyond the three active site motifs. The class B β-lactamases are zinc metalloenzymes and are completely distinct from the serine β-lactamases in terms of structure, mechanism, and evolutionary heritage (Hall et al., 2004). There are three subclasses of class B metallo-β-lactamases (B1 to B3). Classes B1 and B3 are able to bind one or two zinc ions (Heinz and Adolph, 2004), whereas the class B2 enzymes appear to be mononuclear (Garau et al., 2005). In the binuclear metalloβ-lactamases, the zinc ions are proximal to each other and are separated by a bridging hydroxide that has been proposed to be the attacking nucleophile in β-lactam hydrolysis. The class B1 and B3 metallo-β-lactamases can also function as mononuclear enzymes, in which a single zinc ion (that occupies the Zn1 site) coordinates the nucleophilic hydroxide; this mechanism has been proposed to predominate in the presence of substrate under physiological conditions (Wommer et al., 2002). The catalytic mechanism proposed for the class B2 metallo-β-lactamases (Figure 1.7) differs from the class B1 and B3 mononuclear mechanisms in that the zinc ion only occupies the second site (i.e. Zn2), a general base activates the nucleophilic water, and the zinc forms a bond with the amine nitrogen of the hydrolyzed β-lactam amide (Garau et al., 2005). 19  The β-lactamases are ancient enzymes (Hall and Barlow, 2004; Hall et al., 2004) that are thought to have been relatively rare until β-lactam antibiotics were introduced into the clinic half a century ago (Palumbi, 2001). The widespread use of carbapenems, the monobactam aztreonam, cephamycins and oxyimino-cephalosporins in the past few decades has apparently led to the evolution of a new generation of β-lactamases, which have an extended substrate spectrum (i.e. extended-spectrum β-lactamases or ESBLs), as well as the development of novel carbapenemases and plasmid-mediated AmpC β-lactamases [for recent reviews, see (Fisher et al., 2005; Jacoby and Munoz-Price, 2005; Poole, 2004)]. Common ESBLs include varieties from the class A β-lactamases TEM, SHV and CTX-M and the class D β-lactamase OXA. These enzymes are typified by a broad substrate spectrum that includes oxyimino-cephalosporins, aztreonam and in some cases, cefepime (Jacoby and Munoz-Price, 2005). Amino acid substitutions responsible for the ESBL phenotype appear to open the β-lactamase active site to permit access to oxyimino-cephalosporins, which often increases the susceptibility of these enzymes to β-lactamase inhibitors, such as clavulanic acid (Sirot et al., 1997). Substitutions which increase active site flexibility can also impart enhanced β-lactam activity at the expense of protein stability, as in the case of the ceftazidimase activity of CTX-M enzymes (Chen et al., 2005). Carbapenemases are derived from classes A, B and D and are a source of considerable concern as they confer resistance to carbapenems in addition to oxyimino-cephalosporins and cephamycins (Nordmann and Poirel, 2002). The class C AmpC β-lactamases provide resistance to cephamycins and oxyimino-cephalosporins and unlike the ESBLs are resistant to inhibition by clavulanic acid (Jacoby and Munoz-Price, 2005). As more than 20 different AmpC enzymes are known to  20  H118  D120  (a)  C221  H263  (b)  (c) Figure 1.7:  Representative class B β-lactamase CphA. (a) Structure of the CphA fold, shown  with a single zinc atom and bound by the substrate biapenem (PDB code 1X8). (b) Close-up view of the CphA active site with several active site residues indicated as sticks, colored by atom type (C atoms, yellow; O, red; N, blue; and S, orange). Biapenem is displayed with purple carbon atoms to distinguish it from the protein carbons. The catalytic zinc and water are shown as grey and cyan spheres, respectively. Hydrogen bonds are represented by green dashes. (c) Proposed mechanism for the enzymatic hydrolysis of biapenem based on the crystal structure of CphA [catalytic scheme reproduced from (Garau et al., 2005)].  21  be encoded on plasmids, these β-lactamases can be readily donated to enhance the resistance profile of bacterial pathogens (Philippon et al., 2002). Resistant PBPs. Many Gram-positive pathogens rely on the expression of resistant PBPs that display unusually low affinities for β-lactam antibiotics. This is because β-lactamase-mediated resistance in Gram-positive cocci is found almost exclusively in staphylococcal species, which carry predominantly narrow-spectrum β-lactamases that exhibit relatively poor activity against semisynthetic penicillins, cephalosporins, and carbapenems.  There are several PBP-mediated mechanisms of β-lactam resistance,  including: acquisition of a ‘new’ less-sensitive enzyme, mutation of an endogenous PBP to weaken β-lactam-inactivation (while retaining a sufficient degree of transpeptidase activity), or upregulation of PBP expression (Fisher et al., 2005).  Important insights into these  phenomena were recently deduced from the crystal structures of several resistant class B HMM PBPs—PBP2a from methicillin-resistant S. aureus (MRSA) (Lim and Strynadka, 2002), PBP2x from the penicillin-resistant Streptococcus pneumoniae (PRSP) (Chesnel et al., 2003; Dessen et al., 2001; Pernot et al., 2004), and PBP5fm from the naturally resistant Enterococcus faecium (Sauvage et al., 2002). PBP2 is essential in S. aureus as it is the only class A HMM PBP among the four PBPs typically present in these bacteria (Massova and Mobashery, 1998). However, MRSA strains exhibiting high-level resistance to methicillin and other clinically important β-lactams have been found to express a fifth PBP, a class B HMM PBP called PBP2a. PBP2a of MRSA has a low affinity for β-lactams and appears to substitute for the β-lactam-inactivated transpeptidase activity of PBP2. Thus, when challenged with β-lactam antibiotics, MRSA will utilize the transglycosylase activity of PBP2 and the transpeptidase functionality of 22  PBP2a to cooperatively synthesize the cell wall. Kinetic measurements of PBP2a indicated that the resistance of this enzyme to β-lactams predominantly derives from its inefficient formation of the acyl-PBP intermediate during catalysis (Fuda et al., 2004; Lu et al., 1999). This was confirmed by the structure of PBP2a, which revealed a distorted active site that requires an energetically costly conformational change (involving α-helix 2 and β-strand 3) for β-lactam acylation (Lim and Strynadka, 2002). Consistent with these observations, reduced transpeptidase activity has been indicated in PBP2a (Graves-Woodward and Pratt, 1998), which suggests that the evolution of PBP2a proceeded to a point of delicate compromise between β-lactam resistance and transpeptidation in order to maintain cell wall integrity in the face of β-lactam exposure. Whereas the acquisition of PBP2a-mediated resistance in MRSA is thought to have been acquired from horizontal gene transfer (Wielders et al., 2001), resistance can also evolve in endogenous PBPs either from the accumulation of point mutations or by recombination with genes from resistant species to produce “mosaic” genes containing blocks of exogenous DNA (Smith et al., 1991; Spratt, 1988). The mutation of PBP2x in pencillin-resistant Streptococcus pneumoniae (PRSP) has been studied extensively and occurs in a mosaic pattern. Strain sp328 harbors the most clinically important mutation, T338A, which was found to result in the loss of an important active site water molecule, weakening the hydrogen bonding network that stabilizes the acyl-PBP complex (Dessen et al., 2001). The S389L and N514H mutations that are also present in this strain were found to sterically hinder favorable interactions with the β-lactam, reducing the acylation rate. An additional mutation, M339F, confers further resistance to strains that possess the T338A mutation. The structure of this variant was found to re-orientate the S337 nucleophile, 23  lowering the reaction rate by 4 to 10-fold (Chesnel et al., 2003). PBP2x from the PRSP strain sp5259 possesses a Q552E mutation which introduces a negative charge near the edge of the active site, which may disfavor interaction with negatively charged β-lactams (Pernot et al., 2004). Other mutations in this strain act in a similar way to residues from PBP2a, altering the conformation of β-strand 3 so that an active site rearrangement is required for acylation. The endogenous β-lactam resistance conferred by PBP5fm of E. faecium is not immediately apparent, although two features have been implicated from structural studies (Sauvage et al., 2002): (1) a rigid and less accessible active site cleft and (2) charge repulsion with the β-lactam carboxylate group due to an active site glutamate (equivalent to Q552E in sp5259 PBP2x).  E. faecium can demonstrate increased β-lactam resistance either by  overexpression or mutation of PBP5fm, such as insertion of a serine residue at position 466 in an active site loop involved in acylation (Rice et al., 2004).  Multidrug efflux pumps.  With the exception of some strains of the  Streptococci, Enterococci and Staphylococci ‘superbugs’, Gram-negative bacteria are generally more resistant to antibiotics and chemotherapeutic agents than Gram-positive bacteria.  This greater intrinsic resistance results from a synergy of two factors, slow  penetration of the outer membrane and active drug efflux (Nikaido, 2001). Antibiotics diffuse slowly across the outer membrane of Gram-negative bacteria due to the specificity and availability of porin channels (for hydrophilic molecules) and the low fluidity of the lipopolysaccharide leaflet (for hydrophobic molecules). Inevitably, however, antibiotics do trickle through the outer membrane and thus an efflux system is required to clear the periplasm of antibiotics. In the case of β-lactam antibiotics, these multidrug efflux systems 24  (Figure 1.5) consist of an inner membrane drug-proton antiporter of the ResistanceNodulation-Division (RND) family, an outer membrane channel protein, and a periplasmic membrane fusion protein adapter that is thought to couple the inner and outer membrane components for direct extrusion of antibiotics across both membranes. Recent years have seen a tremendous increase in our understanding of the structural and mechanistic features of these tripartite efflux pumps due to crystal structures of all three components, including: antiporter [AcrB from E. coli (Murakami et al., 2002; Yu et al., 2003)], channel [TolC from E. coli (Koronakis et al., 2000)], and adapter [MexA from Pseudomonas aeruginosa (Akama et al., 2004; Higgins et al., 2004) and AcrA from E. coli (Mikolosko et al., 2006)]. Studies of the protein-protein interactions between the three components of the AcrAB-TolC efflux system from E. coli suggest that the periplasmic adapter is essential to forming a stable antiporter-channel complex (Touze et al., 2004) and has allowed the construction of models of the fully assembled pump complex (Eswaran et al., 2004). The protomers of the trimeric antiporter AcrB have been captured in three distinct conformations corresponding to states of drug ‘access’, ‘binding’ and ‘extrusion’ (Murakami et al., 2006; Seeger et al., 2006). These structures suggest a unique three-step rotating ‘peristaltic pump’ mechanism of drug export powered by proton-motive force. Export is initiated by a drug molecule entering the periplasmic vestibule of an AcrB protomer in the ‘access’ state. In the ‘binding’ state, a proton is transported to the cytoplasm and the verstibule opens into a voluminous aromatic binding cavity capable of accommodating a variety of compounds. A new periplasmic proton is bound in the ‘extrusion’ state in concert with a shrinking of the drug-binding cavity. This pushes the drug towards the channel component TolC, which ‘twists’ open due to an allosteric conformational change mediated  25  by the adapter AcrA (Lobedanz et al., 2007). Assembly of these tripartite efflux systems from their components appears to be constitutive (Touze et al., 2004), a feature which may contribute to the rapid extrusion of β-lactams and other compounds [approximated at hundreds of molecules per second (Narita et al., 2003)]. Multidrug efflux is emerging as an important determinant of antibiotic resistance in the Gram-negative pathogen P. aeruginosa (Livermore, 2002), which is predicted to possess eleven such efflux systems (Poole, 2005). The best-characterized of these is MexAB–OprM (adapter, MexA; antiporter, MexB; and channel, OprM). The MexAB-OprM system acts on a wide range of antibiotics—including tetracycline, chloramphenicol, quinilones, novobiocin, macrolides and trimethoprim, as well as β-lactams—and has been directly implicated in clinical resistance to the penicillin ticarcillin (Boutoille et al., 2004; Cavallo et al., 2002) and the carbapenem meropenem (El Amin et al., 2005; Pournaras et al., 2005).  Increased  resistance in these cases appears to have resulted from hyper-production of MexAB-OprM via mutation of its cognate transcriptional regulators (Cao et al., 2004; Poole et al., 1996; Sobel et al., 2005). Expression of mexAB-oprM is primarily regulated by the MarR family repressor, MexR (Saito et al., 2001), but the signals that influence MexR are currently unknown. A defining feature of the MarR family is their common affinity for small phenolic anions (Wilkinson and Grove, 2006). These ligands are typically efflux substrates and appear to function as allosteric effectors that modulate DNA-binding in their cognate regulators. While no such compounds have been definitively identified for MexR, the structure of MexR in the absence of ligands suggests that MexR may uniquely bind a peptide or protein effector (Lim et al., 2002). Like other MarR members, MexR binds its pseudo-palindromic DNA  26  operators (Evans et al., 2001) as a homodimer using two winged-helix DNA-binding domains. The spacing between these two DNA-binding domains can vary considerably as evidenced by the four distinct conformations captured in the MexR crystal structure (Lim et al., 2002). This conformational flexibility is expected to play an important role in the allosteric control of MexR-mediated repression of mexAB-oprM. A number of MarR family proteins have been structurally characterized in recent years, but only a few of these structures have included bound ligands (Alekshun et al., 2001; Hong et al., 2005; Newberry et al., 2007; Saridakis et al., 2008). These structures have identified several distinct effectorbinding sites, but it is presently unclear if any of these correspond to a putative ligand binding site within MexR. Other β-lactam resistance mechanisms. In addition to the active efflux of β-lactams, the deletion of porins has been shown to contribute to increased resistance in a variety of Gram-negative species by further restricting the access of these drugs to their periplasmic targets (Fisher et al., 2005). While the relative importance of this resistance determinant is uncertain, porin deletion can contribute significant resistance to β-lactams when coupled with the expression of ESBLs, as observed in the important pathogen Klebsiella pneumonaie (Bradford et al., 1997; Cao et al., 2000; Nelson et al., 2003). Brief exposure to β-lactams induces the expression of over 200 genes in S. aureus (Utaida et al., 2003). A number of these upregulated genes have clear roles in building and maintaining the cell wall, but a great many have undefined functions. About half of the genes induced by β-lactams are also induced by non-β-lactam inhibitors of cell wall biosynthesis, suggesting the possibility of a co-ordinately regulated ‘stimulon’ that responds to cell wall stress. A particularly interesting consequence of β-lactam exposure is induction 27  of the ‘SOS response’ (Maiques et al., 2006; Miller et al., 2004), an error-prone DNA repair system activated by DNA damage or environmental stress. The SOS response has a variety of diverse triggers such as UV-light, organic mutagens, quinolones, physical stress, and β-lactams (Kelley, 2006). The genes involved in the SOS response can be few or many [for example, 15 genes in P. aeruginosa (Cirz et al., 2006) and 63 genes in Bacillus subtilus (Goranov et al., 2006)]. SOS genes are controlled by the negative and positive regulators, LexA and RecA, respectively, and encode proteins involved in such functions as postreplication DNA repair, recombination, and cell division. Regardless of the stress signal, the formation of single-stranded DNA (ssDNA) appears to ultimately be responsible for triggering the SOS response by interacting with RecA to form nucleoprotein filaments. These ssDNA-RecA filaments assist in the autoproteolysis of the SOS gene repressor LexA (Luo et al., 2001; VanLoock et al., 2003). This cleavage inactivates LexA repression and permits transcription of the SOS genes. One of the proteins encoded by these genes is SulA, which prevents the polymerization of FtsZ (Cordell et al., 2003), a process that is essential in formation of the septum during cell division. Because β-lactam antibiotics only kill dividing bacteria, arresting cell division during β-lactam exposure provides temporary protection against β-lactam lethality. Considering that the SOS response encourages mutation (Cirz et al., 2005) as well as facilitates horizontal gene transfer (Maiques et al., 2006), the induction of the SOS response by β-lactams constitutes both a novel resistance mechanism and a mode of enhancing the evolution of resistance.  28  1.4.3  Regulation of β-Lactam Resistance. Resistance genes can be expressed constitutively or in response to environmental  signals, such as β-lactam exposure. A number of two-component regulation systems are associated with β-lactam-inducible resistance, including 13 out of 32 such systems in E. coli (Hirakawa et al., 2003). Two-component systems communicate environmental cues via phosphorylation (or methylation) signaling pathways; they are comprised of a membranebound sensor histidine kinase and a cytoplasmic response regulator (Mascher et al., 2006). The genes upregulated by β-lactam exposure in two-component systems correspond to various multidrug efflux pumps and β-lactamases, but many of their cognate ligands/signals remain uncharacterized. For example, the two-component system, BlrAB, in Aeromonas spp. has been shown to regulate three β-lactamase genes corresponding to classes B, C and D, but it is presently unknown how the presence of β-lactams activates the sensor histidine kinase, BlrB (Avison et al., 2004). Many Gram-negative bacteria express β-lactamases (including classes A, B and C) under the β-lactam-inducible control of the LysR family regulator, AmpR (Lindberg and Normark, 1987; Okazaki and Avison, 2008). The best characterized example is for the chromosomal class C β-lactamase AmpC. In the absence of β-lactams, AmpR is modulated by association with the peptidoglycan precursor, UDP-MurNAc-pentapeptide, and acts as a repressor of ampC transcription (Jacobs et al., 1997). In the presence of β-lactams, there is an increased degradation of cell wall-derived muropeptides (Jacobs et al., 1994). This leads to the accumulation of anhydro-MurNAc-tripeptide in the cytoplasm, which displaces UDPMurNAc-pentapeptide from AmpR and converts AmpR into an activator of ampC transcription (Jacobs et al., 1997). The result of this tug-o’-war between two ligands is a 29  highly sensitive defense mechanism against β-lactam-induced cell wall damage. Other LysR family regulators, such as SmeR and NmcR, act as weak activators of β-lactamase expression in the absence of β-lactams and stronger activators in their presence (Naas et al., 1995; Naas and Nordmann, 1994). Similar LysR regulators have been found controlling β-lactamase expression in the Gram positive bacterium Streptomyces cacaoi (Magdalena et al., 1997). A number of Gram positive bacteria regulate the production of class A β-lactamases by an uncommon proteolytic transmembrane signaling pathway that is inducible by β-lactam antibiotics (Zhang et al., 2001). The two key regulators in the S. aureus system are the transmembrane sensor/transducer BlaR1 (a class C HMM PBP) and the repressor BlaI, which prevents transcription of the β-lactamase gene blaZ (Figure 1.5). When an extracellular β-lactam binds to the transpeptidase-like ‘sensor’ domain of BlaR1, the zymogenic ‘transducer’ domain is activated by proteolytic cleavage on the cytoplasmic side of the membrane. This activated BlaR1 protease then induces transcription of blaZ by destroying the affinity of BlaI for its cognate DNA operator through cleavage of its dimerization domain. Intriguingly, PBP2a production in MRSA is regulated by a β-lactam-inducible system that is homologous to the BlaI/BlaR system, including a repressor MecI and sensor/transducer MecR that binds β-lactams (Zhang et al., 2001). Many β-lactam resistance genes are not inducible by β-lactams at all, but are instead expressed constitutively at basal (i.e. relatively low) levels. For example, production of the prominent MexAB-OprM multidrug efflux pump in P. aeruginosa is regulated by the MarR family repressor, MexR (Saito et al., 2001). While the signals that modulate MexR are currently unknown, a defining feature of the MarR family is their common affinity for small phenolic anions (Wilkinson and Grove, 2006). These ligands are typically efflux substrates 30  and appear to function as allosteric effectors that modulate DNA-binding in their cognate regulators.  While no such compounds have been definitively identified for MexR, the  structure of MexR in the absence of ligands suggests that MexR may uniquely bind a peptide or protein effector (Lim et al., 2002). Like other MarR members, MexR binds its pseudopalindromic DNA operators (Evans et al., 2001) as a homodimer using two winged-helix DNA-binding domains. The spacing between these two DNA-binding domains can vary considerably as evidenced by the four distinct conformations captured in the MexR crystal structure (Lim et al., 2002). This conformational flexibility is expected to play an important role in the allosteric control of MexR-mediated repression of mexAB-oprM. A number of MarR family proteins have been structurally characterized in recent years, but only a few of these structures have included bound ligands (Alekshun et al., 2001; Hong et al., 2005; Newberry et al., 2007; Saridakis et al., 2008). These structures have identified several distinct effector-binding sites, but it is presently unclear if any of these correspond to a putative ligand binding site within MexR. Hyperexpression of resistance genes due to the mutation of regulatory elements (constitutive or inducible) is a common route to increased resistance in the presence of antibiotic pressure. While these mutations are often associated with a fitness cost in the absence of antibiotics (Folkesson et al., 2005; Katayama et al., 2003; Sanchez et al., 2002), additional mutations can sometimes restore fitness without compromising antibiotic resistance (Bjorkman et al., 2000).  31  1.5  OBJECTIVES OF THESIS The continued evolution and dissemination of antibiotic resistance in pathogenic  bacteria demands a sustained stream of innovations in antimicrobial therapy to avert the emergence of panresistant superbugs.  Furthermore, the diminishing interest of the  pharmaceutical sector in the development of novel antibiotics implies that government and academic labs will share the greatest degree of responsibility in elucidating the molecular details of resistance. This thesis aims to contribute to our understanding of how resistance to β-lactam antibiotics is regulated in two notoriously resistant bacterial pathogens.  It is  heartening that bacteria regulate resistance mechanisms at all, as it suggests that—at least in some cases—constitutive expression of resistance determinants incurs a fitness cost that weakens their survival in the absence of antibiotic pressure. The common theme of these studies is the structural characterization of bacterial proteins involved in switching ‘on’ β-lactam resistance. Chapters 2 and 3 focus on the ‘sensing’ of β-lactams by the BlaR1 sensor/transducer, a receptor involved in the control of β-lactamase production in S. aureus. In Chapter 2, the first crystal structures of the S. aureus BlaR1 sensor domain are described, both free of substrate and bound by penicillin. Mechanistic insights into the acylation of this domain by β-lactams are also discussed. Chapter 3 probes these BlaR1 sensor domain structures for clues regarding the signal transduction mechanism and provides support for the hypothesis that an extracellular loop of BlaR1 plays an important role in transmitting this signal. Chapter 4 investigates a putative 53-residue protein effector called ArmR involved in regulating expression of the prominent multidrug efflux pump MexAB-OprM from P. aeruginosa. In-vitro studies are described that confirm that ArmR functions to neutralize the DNA-affinity of the MarR family 32  repressor, MexR. Following this is a description of the crystal structure of MexR double mutant Q106L/A110L (MexRLL) in complex with ArmR residues 29-53 (ArmRC), the first structure of a MarR protein bound by a protein effector. Comparison of this structure to previously solved structures for apo MexR reveals the allosteric mechanism for alleviating repression of MexAB-OprM-mediated efflux.  33  CHAPTER 2 – The β-Lactam Sensor of BlaR1  2.1  INTRODUCTION Resistance to penicillin was first reported in a strain of Staphylococcus aureus in  1944, only a few years after it was first introduced into the clinic (Kirby, 1944). Today, only a few percent of S. aureus infections are susceptible to penicillin as more than 95% of clinical isolates express β-lactamases (Lowy, 1998) and approximately 60% of isolates express PBP2a (NNIS, 2004), which confers broad-spectrum insensitivity to β-lactam antibiotics.  Until very recently, the glycopeptide vancomycin was the only effective  treatment for serious MRSA infections, but reports of intermediate (CDC, 2000; Hiramatsu et al., 1997; Sieradzki et al., 1999; Smith et al., 1999) and full resistance (CDC, 2004; Chang et al., 2003; Weigel et al., 2003) to vancomycin have emerged in the past decade. Several new agents with anti-MRSA activity have been introduced in the last few years (quinupristindalfopristin, linezolid, daptomycin, and tigecycline) (Rice, 2006), but resistance to all of these drugs except tigecycline has already been reported in Enterococcus faecalis isolates (Chow et al., 1997; Gonzales et al., 2001; Lewis et al., 2005), foreshadowing the acquisition of resistance in strains of S. aureus in the near future. S. aureus is the principal cause of both community-acquired and nosocomial bacteremia, particularly in intensive care units where it is a leading cause of death with mortality rates of 20-50% (Blot et al., 2002; Cheong et al., 1996; Selvey et al., 2000). Moreover, the economic burden of fighting these infections is enormous, annually costing Canadians $100’s of millions (Kim et al., 2001) and billions in the United States (Gould, 2006).  34  Expression of the structural genes for β-lactamases and PBP2a (i.e. blaZ and mecA, respectively) are regulated by orthologous systems (Figure 2.1) consisting of a repressor (BlaI/MecI) and a sensor/transducer protein (BlaR1/MecR1) that destroys the activity of the repressor when activated by antibiotics. The BlaI and MecI repressors form homodimers that specifically bind DNA operator/promoter sequences. BlaR1 and MecR1 are membranespanning multidomain proteins, each composed of three domains: (i) an extracellular C-terminal β-lactam sensor domain, (ii) an N-terminal transmembrane domain containing four predicted transmembrane α-helices (Hardt et al., 1997), and (iii) an intracellular metalloprotease domain that is thought to proteolytically inactivate the BlaI/MecI repressor in the cytoplasm. Existing evidence suggests that binding of β-lactam antibiotics to the sensor domain causes a conformational change that results in autolytic activation of the intracellular protease domain (Zhang et al., 2001), allowing it in turn to catalyze cleavage of BlaI/MecI (directly or indirectly) at a site critical for dimerization. This cleavage event effectively destroys the ability of the repressor to bind DNA, permitting the transcription of not only blaZ/mecA, but blaI/mecI and blaR1/mecR1 as well. In this system, expression of these proteins is efficiently terminated when the signaling antibiotic levels are reduced (primarily through hydrolysis by surrounding β-lactamase enzymes (Zhang et al., 2001)). Whereas the bla divergeon (i.e. blaZ, blaI, and blaR1) is plasmid-borne, constitutive β-lactamase expression has been observed in S. aureus strains possessing normal penicillinase plasmids (Cohen and Sweeney, 1968). Several explanations for this observation have been proposed, including the involvement of an as of yet unidentified chromosomally encoded regulatory component known as BlaR2 (Filee et al., 2002; Garcia-Castellanos et al., 2004). 35  BlaR1 and MecR1 from S. aureus share significant sequence identity (sensor domains, 43%; protease domains, 33%; and full-length proteins, 34%), and the regulatory genes of bla and mec have been shown to be interchangeable in vivo although each sensor/transducer is specific only for its corresponding repressor (Lewis and Dyke, 2000; McKinney et al., 2001). Indeed, due to the potency of the MecI repressor, many methicillinresistant S. aureus (MRSA) strains have MecI deletions that inactivate repression (Suzuki et al., 1993), incurring either constitutive expression of PBP2a or inducible expression regulated by BlaR1/BlaI. The absence of the bla or mec regulatory genes has been observed to select against PBP2a expression, implicating a role for these genes in stabilizing dissemination of mecA to new host strains (Katayama et al., 2003). Crystal structures have been determined for the S. aureus repressors BlaI (Safo et al., 2005) and MecI both in their apo forms (Garcia-Castellanos et al., 2003) and in complex with DNA (Garcia-Castellanos et al., 2004; Safo et al., 2006). These repressors display significant flexibility which permits BlaI to bind the mec operator, MecI to bind the bla operator and suggests that repressor proteolysis occurs while DNA-bound.  In contrast, the BlaR1 and  MecR1 sensor/transducers have proved a much greater challenge for structural biologists. Early sequence comparisons indicated that the sensor domain adopts a fold that closely resembles the class D β-lactamaes (Zhu et al., 1990). This was confirmed by the crystal structure of the BlaR sensor domain from Bacillus licheniformis, solved in the absence of acylating β-lactam (Kerff et al., 2003). In contrast to the class D β-lactamases, however, was the lack of Nζ-carboxylation at a critical active site lysine.  This lysine residue  (corresponding to the SXXK motif found in all penicilloyl serine hydrolases) was previously shown to undergo carboxylation in the BlaR1 sensor domain of S. aureus (Golemi-Kotra et  36  al., 2003). This chapter describes the first crystal structures of the S. aureus BlaR1 β-lactam sensor domain (hereafter referred to as BlaRS) in both its apo and penicillin-acylated forms. These structures illuminate the active-site features that are responsible for the PBP activity of BlaRS and provide mechanistic insights into the role of BlaRS in detecting β-lactam antibiotics and transducing the binding signal across the bacterial cell membrane.  Figure 2.1: Regulation systems controlling the expression of β-lactamase and PBP2a (shown in large brackets). Genes and the proteins they encode are matched by color. S  Dotted arrows represent gene  P  expression. BlaR = BlaR1 sensor domain. BlaR = BlaR1 protease domain. 3D structures were derived from published atomic coordinates (BlaI/MecI – PDB ID 1OKR, β-lactamase – PDB ID 1M6K, PBP2a – PDB ID 1MWR).  37  2.2  METHODS  2.2.1  Cloning, Protein Expression, and Purification BlaR1 was cloned from a plasmid containing the bla divergeon (p184R6H) as  described previously (Zhang et al., 2001). This plasmid was originally derived from a β-lactamase expressing strain of S. aureus (67-0) isolated from a patient at San Francisco General Hospital Medical Center (Chambers et al., 1985; Hackbarth and Chambers, 1993). The BlaR1 sensor domain (amino acids 330–585, i.e. BlaRS) was subcloned into the pET41b(+) vector (Novagen) and transformed into Escherichia coli Rosetta (DE3) (Novagen). Cells were grown from an overnight culture at 37 °C until A600 ~ 0.6, heatshocked at 42 °C for 1 h, cooled to 15 °C, and then induced to overexpress BlaRS overnight using 1 mM isopropyl-β-D-thiogalactopyranoside (IPTG). This same expression procedure was adapted for the production of selenomethionine (SeMet)-substituted BlaRS using previously described protocols (Doublie, 1997). The cells were lysed by high pressure homogenization using an Avestin EmulsiFlex-C5. The BlaRS protein was purified from the soluble cell fraction using three chromatographic steps performed at 4 °C. The soluble cell lysate was first bound to Fractogel EMD SO3– resin (Novagen), pre-equilibrated in 20 mM HEPES (pH 7.5) and 50 mM NaCl, and eluted with 0.8 M NaCl. The eluate was dialyzed overnight at 4 °C in buffer containing 0.2 M NaCl and then filtered using 0.22-µm filters. The filtrate was further purified using a Mono S HR 5/5 cation exchange column (Amersham Biosciences) and a 0.2–0.8 M NaCl elution gradient. Elution fractions corresponding to the principal A280 peak were combined, concentrated, and passed through a Superdex 200 HR 10/30 size exclusion column (Amersham Biosciences) equilibrated in 20 mM Tris-HCl (pH 7.5) and 150 mM NaCl as the storage buffer. The single A280 peak was collected and 38  concentrated to 20–40 mg/ml as estimated using a predicted 280 nm molar absorption coefficient of 60280 M–1 cm–1 (Pace et al., 1995). Concentrated BlaRS was either used fresh (stored at 4 °C) or flash-frozen in liquid N2 and stored at -80 °C. 2.2.2  Crystallization and Data Collection Crystals of apo- and acyl-BlaRS were obtained using the hanging drop vapor diffusion  method. For apo-BlaRS crystals, 1 µl of a 20 mg/ml protein solution was added to equal volumes of reservoir solution consisting of 18–23% polyethylene glycol 3350 and 0.2 M NaH2PO4. Equilibration over a 0.5-ml reservoir at 18 °C for 1–2 weeks produced single crystals reaching dimensions of up to 0.7 × 0.5 × 0.4 mm. For acyl-BlaRS, benzylpenicillin (PenG) was incubated with BlaRS at final concentrations of 20 mg/ml protein and 10 mM PenG for 15 min at room temperature before initiating crystallization. Native acyl-BlaRS crystals were then grown using 1 µl of this protein solution mixed with an equal volume of reservoir solution consisting of 26–29% polyethylene glycol 3350, 0.2 M NaCl, and 0.1 M BisTris (pH 6.6). Crystals were typically observed after 1 week at 18 °C. SeMet crystals of acyl-BlaRS were obtained by streak-seeding native crystals into fresh drops containing SeMet-substituted BlaRS, benzylpenicillin, and crystallization reagents at the same concentrations used to grow the native crystals. Crystal clusters formed in 3–5 days with maximum dimensions of 0.4 × 0.2 × 0.15 mm for the individual crystals of the cluster. Single SeMet acyl-BlaRS crystals were obtained by gently breaking the clusters. Diffraction data were collected at 100 K using cryoprotectant solutions of 35% polyethylene glycol 3350 and 0.3 M NaH2PO4 for the apo-BlaRS crystal and 35% polyethylene glycol 3350, 0.5 M NaCl, and 0.1 M BisTris (pH 6.6) for the SeMet acyl-BlaRS crystal. All data sets were collected at the National Synchrotron Light Source on beamline X8-C using an ADSC Quantum Q4R  39  CCD detector. Data were processed using the HKL package (Otwinowski and Minor, 1997) and programs from the CCP4 software suite (Collaborative Computational Project, 1994). The apo crystal was of space group P21, with four molecules/asymmetric unit and unit cell dimensions of a = 59.9, b = 104.9, and c = 90.3 Å and β = 107.7°. SeMet acyl-BlaRS crystallized in space group P43212, with two molecules/asymmetric unit and unit cell dimensions of a = b = 88.4 and c = 125.1 Å. Statistics for data collection and processing are summarized in Table I. 2.2.3  Structure Solution and Refinement The structure of acyl-BlaRS was determined using single anomalous dispersion with  the peak data to locate the initial selenium sites in SOLVE (Terwilliger and Berendzen, 1999), followed by three wavelength multiple anomalous dispersion to generate more accurate phases. Although 100% incorporation of SeMet into BlaRS was indicated by mass spectrometry, only 15 of 18 possible selenium sites were located. Phases were improved with density modification using RESOLVE (Terwilliger, 2000). The initial model was automatically built with RESOLVE (50% complete) (Terwilliger, 2002) and manually rebuilt using XTALVIEW (McRee, 1999). Iterations of refinement with CNS (Brunger et al., 1998) and REFMAC (Collaborative Computational Project, 1994) resulted in the final model. Of the 255 residues in the BlaRS construct, 242 residues were modeled in chain A (residues 4-13, 18-201, and 204-251), and 241 residues were modeled in chain B (residues 8-26 and 31-252). Model quality was analyzed using PROCHECK (84% in the most favorable region of the Ramachandran plot) (Laskowski et al., 1993). Asn388 is a well ordered active-site residue, but adopts a disallowed main chain conformation due to its juxtaposition to benzylpenicillin-acylated Ser389. The apo-BlaRS structure was solved by molecular  40  replacement using the acyl-BlaRS structure as a starting model and the program Molrep (Vagin, 1997). Model rebuilding was performed with XTALVIEW using a prime-and-switch map generated by RESOLVE to reduce model bias. The model was refined and analyzed for quality as described above. The resulting model consisted of 244 residues for chain A (residues 8–166 and 169–253), 247 residues for chain B (residues 8–254), 243 residues for chain C (residues 5–13, 15–76, 78–84, 86–202, and 206–253), and 240 residues for chain D (residues 7–76, 80–82, 84–166, and 169–252). In each of the four molecules of the apo form, a pyrophosphate was also modeled into the active site. This ligand is only 50% occupied in three of the four molecules of the asymmetric unit, as revealed by the close proximity of a partially occupied water molecule to one of the phosphorus atoms. Considering that the crystallization conditions for apo-BlaRS contained 0.2 M sodium phosphate, the unknown electron density could also be modeled as two partially occupied phosphate ions, but the staggered geometry of the tetrahedral phosphorus atoms and the bent P–O–P bond indicate a molecule of pyrophosphate, introduced as a sodium phosphate impurity. The final apo-BlaRS model had 88% of the residues in the most favorable region of the Ramachandran plot and none in the disallowed region. The multiple anomalous dispersion phasing and model refinement statistics for both structures are provided in Table I. Fig. 3 was prepared with MOLSCRIPT (Kraulis, 1991) and rendered with Raster3D (Merritt and Bacon, 1997). All other protein graphics (Figs. 5–7) were prepared and rendered with PyMOL (DeLano, 2002). 2.2.4  Static Light Scattering Static light scattering experiments were performed at 25 °C on a Superdex 75 HR  10/30 size exclusion column (Amersham Biosciences) using 50 mM HEPES (pH 7.5) and 100 mM NaCl. Protein concentration was 1 mg/ml. Refractive index and miniDAWN light  41  scattering detectors (Wyatt Technology Corp.) were calibrated using bovine serum albumin (Sigma). 2.2.5  Mass Spectrometry Apo- and acyl-BlaRS samples were prepared by briefly incubating the pure BlaRS  protein at room temperature in the presence and absence of 10 mM PenG. These samples were then injected onto an ultrafast microprotein analyzer (Michrom BioResources, Pleasanton, CA) equipped with a 1 × 50-mm PLRP-S reverse phase column. The protein was subsequently eluted with a 2–90% gradient of acetonitrile in water and 0.05% trifluoroacetic acid at a flow rate of 50 µl/min over 5 min. Mass analysis of the eluted protein was performed using a PE-Sciex API 300 triple-quadrupole mass spectrometer scanned over a mass-to-charge ratio range of 300–2200 Da with a step size of 0.5 Da and a dwell time of 1.5 ms/step. The ion source voltage was set at 5.5 kV, and the orifice energy was 45 V. Protein molecular masses were determined from these data using the deconvolution software supplied by PE-Sciex.  2.3  RESULTS  2.3.1 Functional Characterization of Recombinant BlaRS The C-terminal extracellular sensor of the BlaR1 signal-transducer from S. aureus (BlaRS) was expressed in the cytoplasm of E. coli as a soluble domain, spanning amino acids 331-585 and lacking an N-terminal Met. To verify the activity of our recombinant protein, the ability of BlaRS to be covalently modified by β-lactam antibiotics was demonstrated using mass spectrometric analysis before and after incubation with benzylpenicillin. This experiment yielded a homogenous mass shift of 333 Da, corresponding closely with the 42  expected mass shift of 334 Da for a benzylpenicillin adduct. Similar deletion mutants containing the sensor domain of BlaR1 have been expressed in E. coli previously for both S. aureus (Golemi-Kotra et al., 2003) and Bacillus licheniformis (Duval et al., 2003) and have been shown to fully retain their activities as highly sensitive penicillin-binding proteins. 2.3.2  Overall Fold and Oligomerization State of BlaRS BlaRS was crystallized in the absence and presence of benzylpenicillin in order to  facilitate determination of the 3D structures of the apo- and acyl-BlaRS proteins by X-ray crystallography.  The crystal structure of acyl-BlaRS was solved first using MAD data  collected from a selenomethionine-substituted crystal.  The final model of acyl-BlaRS  consisted of two molecules per asymmetric unit which superposed with a root mean squared deviation (rmsd) of 0.58 Å on the 232 commonly observed Cα atoms. The model was refined to 2.4 Å resolution with R and Rfree values of 22.0% and 27.6%, respectively (Table 2.1). The atomic co-ordinates from the acyl-BlaRS structure were then used to solve the crystal structure of apo-BlaRS by molecular replacement. The resulting model had four molecules in the asymmetric unit and was refined to 1.8 Å resolution with respective R and Rfree values of 18.6% and 22.7% (Table 2.1). The four molecules superpose closely with rmsd values of 0.33-0.43 Å on the 234-244 commonly observed C-α atoms. The main-chain fold of BlaRS consists of two domains, one helical and one mixed α/β with the penicillin-binding site residing at the interface (Figure 2.2). The mixed α/β domain includes a seven-stranded β-sheet, comprised of six antiparallel strands and a short parallel strand, sandwiched between a pair of α-helices on either side. The helical domain is composed of six helices with the active site situated in an inter-domain cleft centered on helix C which consists of a single 310-helical turn followed by three α-helical turns. An 43  Ω-loop connects helices G and H and forms one edge of the active site. There is no obvious aromatic or hydrophobic patch on the surface of BlaRS, and addition of detergent or lipid was not required for solubility or activity, suggesting no close association between the sensor domain of BlaR1 and the bacterial membrane. Table 2.1: Crystallographic data and refinement statistics.  Data collection Apo  Pen G-acylated, SeMet-derivitized Peak  Spacegroup  Inflection  Remote  Hi-Res  P21  P43212  Molecules/A.U.  4  2  Resolution (Å)  50-1.8  20-2.5  20-2.5  20-2.5  50-2  Wavelength (Å)  1.10000  0.97911  0.97940  0.90000  1.10000  Total reflections  380,826  136,340  91,938  72,967  217,169  95,275  32,152  32,032  31,702  33,767  97.3 (78.0)  98.5 (97.5)  98.1 (97.1)  97.2 (97.1)  98.7 (94.4)  27.2 (3.2)  23.0 (6.8)  19.4 (5.4)  18.9 (5.0)  31.4 (3.0)  4.7 (27.2)  4.7 (12.1)  4.4 (12.2)  3.8 (11.2)  5.6 (40.6)  Unique reflections Completeness (%)  a  I / σ(I)a Rsym (%)  a,b  Refinement statistics Resolution (Å)  Apo 50-1.8  Pen G, SeMet 15-2.4  Rcryst/Rfree (%)c  18.6 / 22.7  22.0 / 27.6  Bonds (Å)  0.012  0.018  Angles (°)  1.30  1.70  Protein  23.4  46.3  Pen. G  N/A  36.3  Water  29.2  38.0  R.m.s Deviations  Average B-factor  a) High resolution shell (1.86 Å – 1.80 Å for apo-BlaRS; 2.07 Å – 2.00 Å for acy-BlaRS) statistics are in parentheses b) Rsym =Σ|(Ihkl)−<I>|/Σ(Ihkl), where Ihkl is the integrated intensity of a given reflection. c)  Rcryst=(Σ|Fo–Fc|)/(ΣFo), where Fo and Fc are observed and calculated structure factors. 5 % of total reflections were excluded from the refinement to calculate Rfree.  44  As observed previously for the B. licheniformis BlaR1 sensor domain (Kerff et al., 2003), the S. aureus BlaRS structure resembles that of the class D β-lactamases, with overall rmsd values in the range of 1.17-1.41 Å on 192-200 common Cα residues for the available OXA structures. Overlapping the apo form of the sensor domains of B. licheniformis and S. aureus gives similar rmsd values of 1.23-1.27 Å on 214 common Cα atoms (36% amino acid sequence identity). The closest match of BlaRS and the class D enzymes occurs with OXA-1 from E. coli which shares 28% amino acid sequence identity with BlaRS. This is consistent with the fact that OXA-1 is the only class D β-lactamase that prefers a monomeric state (Sun et al., 2003). Other class D β-lactamases have been shown to exist predominantly as dimers in solution, an oligomeric form which promotes maximal catalytic activity and which can be mediated by ion binding (Paetzel et al., 2000). BlaRS was observed as a monomer in either crystal form as well as in solution as determined by static light scattering at similar protein concentrations (data not shown). Examination of the BlaRS structure indicates it has lost the prominent dimeric interface observed in the class D crystal structures. A comparison of BlaRS with the OXA-10 dimer reveals that many of the residues responsible for stabilizing the OXA dimer are different in BlaRS, precluding the formation of two salt bridges, three hydrophobic interactions, and three metal ligands. Although a pseudo-two-fold symmetry axis is created by crystal packing in both of our structures that loosely resembles the OXA dimer, the surface area buried by the monomer-monomer interface is almost nonexistent, the strands of the intermolecular β-sheet meet at a steep angle of ~30°, and only a total of four hydrogen bonds join the two molecules.  45  Figure 2.2: Structure of apo-BlaRS. Helices are shown in yellow, strands in red. The catalytic Ser389 and Lys392 residues have been shown in ball-and-stick representation.  2.3.3  BlaRS Active Site Architecture and the Benzylpenicillin Adduct Three conserved sequence motifs define the active site of all penicilloyl serine  transferases/hydrolases, including the class A, C, and D β-lactamases, the cell wall transpeptidases, and BlaR1/MecR1 (Figure 2.3). In BlaRS these motifs correspond to Ser389X-X-Lys392 (with Ser389 the proposed nucleophile in acylation), Ser437-X-Asn439, and Lys526Thr527-Gly528 (Figure 2.4). Within the α-helical domain, the SXXK motif lies at the center of the active site cleft at the N-terminal end of helix C. The SXN element is adjacent on a short loop connecting helices E and F and the KTG motif is situated nearby on strand H of the α/β domain. Flanking Lys392 and opposite Ser389 are several hydrophobic residues, including Leu395 and Phe112 as well as Trp145 and Met146 which are positioned on a 46  proximate Ω-loop (Asn465-Lys481).  The putative oxyanion hole is composed of the  backbone amide nitrogens of Ser389 and Thr529. A hydrogen bond network connects the Oδ of Asn439, the Nζ of Lys392, the Oγ of Ser437, and the Nζ of Lys526. From sequence information, it is expected that the other species of BlaR1/MecR1 sensor domain should share highly similar active site features with BlaRS. The only comparison available at the time these results were first published was with the apo form of the sensor domain of B. licheniformis. As predicted, the two active sites are highly similar (overall rmsd on all atoms of common active site residues i.e. S389, K392, S437, F442, W475, K526, T527, and T529 is 0.68-0.70 Å) despite the unique substitution of threonine in the position of Asn439 in the B.licheniformis species (Kerff et al., 2003). An additional feature observed in the active site of our apo-BlaRS is a molecule of partially occupied pyrophosphate which hydrogen bonds with the Oγ of Ser389 (2.4 Å), the Nζ of Lys526 (2.9 Å), the side chains of Thr527 and Thr529, the backbone carbonyl of Thr529, and the amide nitrogens of Ser389 and Thr529 that comprise the oxyanion hole. In the acyl-BlaRS crystal, benzylpenicillin is unambiguously observed as a covalent adduct of Ser389 in both molecules of the asymmetric unit with C7 bound to the Oγ of the proposed nucleophile Ser389 via an ester linkage (Figure 2.5a). The adduct has been refined at full occupancy in each molecule, with average B-factors of 34.8 and 37.8 Å2 (similar to the average B-factor of 31.1 Å2 observed for the surrounding active site residues). The backbone nitrogens of Thr529 and Ser389 form hydrogen bonds with the carbonyl oxygen of the adduct ester, creating the oxyanion hole typical of β-lactamases and serine proteases. The thiazolidine methyls are stabilized by hydrophobic interactions with the side chains of Phe421 and Thr529 while hydrogen bonds with the side chain of Asn439 and the backbone 47  Figure 2.3:  Amino acid sequence alignment of BlaRS with the BlaR1 sensor domains of  Bacillus licheniformis (BlaR1-Bl), Staphylococcus haemolyticus (BlaR1-Sh), the MecR1 sensor domains  of  S.  aureus  (MecR1-Sa),  Staphylococcus  epidermis  (MecR1-Se),  and  a  representative class D β-lactamase OXA-10 from Pseudomonas aeruginosa (OXA10-Pa). Conserved active site residues have been highlighted in yellow while the remaining active site residues have been highlighted in green. Conserved residues in the rest of the protein have been highlighted in magenta. The alignment was performed with T-Coffee (Notredame et al., 2000).  48  Figure 2.4: Active site of (a) apo- and (b) acyl-BlaRS. The carbons belonging to the benzylpenicillin adduct have been shown in magenta. Hydrogen bonds are indicated by dashed yellow lines.  Figure 2.5: Electron density of Lys392 and PenG adduct. (a) Weighted 2Fo-Fc electron density (contoured at 1.0 σ) is shown around the side chain of Lys392 and the PenG adduct. (b) Weighted Fo-Fc omit map (contoured at 2.0 σ) used to model the PenG adduct. The final refined adduct is shown within the electron density.  49  carbonyl of Thr529 position the adduct amide. The thiazolidine carboxylate is fixed by hydrogen bonds with the side chains of Thr527 and Thr529, a feature typical of PBPs. Interestingly the extra arginine residue utilized by the various classes of serine β-lactamases to form electrostatic interactions with the thiazolidine carboxylate of the substrate is absent in the BlaRS structure, a scenario also typical of PBPs. The binding mode of benzylpenicillin is highly similar in both molecules of the asymmetric unit, excepting the less ordered side chain phenyl substituent (Figure 2.5b) which adopts alternate conformations in each case (making hydrophobic interactions with the side chains of Ile531, Thr529, Phe421, and Tyr536 or the side chains of Ile531, Met476, and the Cβ, Cγ of Glu477).  2.4  DISCUSSION  2.4.1  The Role of Lys392 as the General Base in Acylation The active sites of the PBPs and class A, C, and D β-lactamases center on a  conserved serine nucleophile, but the general base involved in catalysis is apparently different in each (e.g. either Lys73 (Golemi-Kotra et al., 2004; Strynadka et al., 1992; Swaren et al., 1995) and/or Glu166 in class A β-lactamases (Lamotte-Brasseur et al., 1991; Minasov et al., 2002; Nukaga et al., 2003), Tyr150 in class C β-lactamases (Oefner et al., 1990), carboxylated Lys70 in class D β-lactamases (Golemi et al., 2001) and an unprotonated Lys392 in the PBPs (Davies et al., 2001; Rhazi et al., 2003) (Figure 2.6). Whichever the class, the role of the general base in acylation is to activate the nucleophile that attacks the β-lactam ring by deprotonation of the catalytic SXXK serine. Deacylation requires either an additional general base to activate water for hydrolysis of the acyl-enzyme intermediate (e.g. class A β-lactamases), or a mechanism for deprotonating/regenerating the first general base 50  (a)  (b)  (c)  (d)  Figure 2.6: Comparison of the apo-BlaRS active site with representative β-lactamases and a PBP.  BlaRS active site residues (tan) are superimposed with the corresponding residues in a  representative (a) class C β-lactamase (AmpC from Escherichia coli, yellow, PDB ID 1KE4), (b) class D β-lactamase (OXA-10 from Pseudomonas aeruginasa, green, PDB ID 1K57), (c) class A β-lactamase (SHV-2 from Klebsiella pneumoniae, purple, PDB ID 1N9B), and (d) PBP (PBP2x from Streptococcus pneumoniae, light blue, PDB ID 1QME). Active site overlays were generated with Swiss-Pdb Viewer (Guex and Peitsch, 1997) by fitting the Cα atoms of Ser389, Lys392, and Asn439 from BlaR1 with the corresponding Cα atoms from the other structures.  51  (e.g. class A, C, and D β-lactamases). In BlaRS, the close proximity of Lys392 (2.4-2.8 Å) and Ser437 (3.2-3.5 Å) to the Ser389 nucleophile suggests two possible candidates for a general base in acylation, the former arguably more suitable in terms of distance and potential pKa to act either in an unprotonated state similar to the PBPs (Kerff et al., 2003) or carboxylated state similar to the class D β-lactamases (Golemi-Kotra et al., 2003). 2.4.2  The Nζ-Carboxylation of Lys392 Carboxylation is favored at basic pH, but has been observed in crystal structures of  the class D β-lactamases as low as pH 6.0 (Golemi et al., 2001) and perhaps even at pH 5.5 with low occupancy (Sun et al., 2003). The structures of apo- and acyl-BlaRS presented here were determined at pH 4.7 and pH 6.6, respectively. In either case, carboxylation of Lys392 was not observed at any contour level in our electron density maps. In the class D βlactamase structures, a non-carboxylated Lys70 (corresponding to Lys392 in BlaR1) seems to encourage an “inactive” conformation of Ser115 (Ser437 in BlaR1) where the serine hydroxyl (presumed to shuttle a proton from the carboxylated lysine to the leaving group nitrogen of the β-lactam substrate) is somewhat displaced from its typical position in the active site (Golemi et al., 2001; Pernot et al., 2001). In contrast with many of the noncarboxylated structures of the class D β-lactamases, Kerff and co-workers noted that the active site of the apo form of the B. licheniformis BlaR sensor domain (which also lacks a carboxylated active site lysine despite the fact that the crystals were grown at pH 7.0) closely resembles the “active” conformation (Kerff et al., 2003). Similarily, we observe the active sites of both apo- and acyl-BlaRS to adopt the “active” conformation in our structures. More recently, this “active” non-carboxylated active site has been observed in the sensor domain structures of ceftazidime-acylated BlaR1 at pH 7.5 (Birck et al., 2004), apo-MecR1 at pH 52  7.0, PenG-acylated MecR1 at pH 6.5 and oxacillin-acylated MecR1 at pH 6.5 (Marrero et al., 2006). To date, no structures of BlaRS or any of its sensor/transducer homologs have been determined showing either a carboxylated Lys392 or an “inactive” active site conformation. These results are surprising since Lys392 of BlaRS has been shown to specifically bind CO2 in solution via  13  C-NMR, incorporation of  14  CO2, fluorescence quenching of  Trp475 (Golemi-Kotra et al., 2003) and Fourier-transformed infrared spectroscopy (FT-IR) (Golemi-Kotra et al., 2003; Thumanu et al., 2006). Moreover, the dissociation constant for BlaRS and CO2 was measured at 0.6 μM (Golemi-Kotra et al., 2003), only a 2- to 3-fold increase versus the class D β-lacatamase OXA-10 (Golemi et al., 2001) and well below 1.3 mM—the physiological concentration of CO2 (Tien et al., 1999). Considering these facts, Lys392 of BlaR1 should be 100% carboxylated in vivo. It should be noted, however, that the carboxylation of Lys392 has not yet been directly shown to be an absolute requirement for the β-lactam-binding activity of BlaR1 in S. aureus.  Moreover, preliminary kinetic  experiments in B. licheniformis suggested no significant increase in acylating activity under conditions which would promote carboxylation of Lys392 in that system (Kerff et al., 2003). With that in mind, the active site of BlaRS differs from class D β-lactamases such as OXA-10 in several interesting ways (Figure 2.6b). The carboxylate of the carboxy-Lys70 of OXA-10 is held in position by hydrogen bonding interactions with Ser67, Trp154, and Asn73 (through a molecule of water) and hydrophobic interactions with Phe120, Ile155, and Val117. Two of these interactions have been disrupted in the BlaRS active site. Strictly conserved in the OXA structures, Val117 has been replaced by Asn439 in the BlaRS structure, which now forms a hydrogen bond with the Nζ of Lys392 (an interaction found in all class A and class C β-lactamases as well as the PBPs, none of which utilize a 53  carboxylated lysine). Likewise, Asn73 from the structure of OXA-10 has been substituted with the aliphatic Leu395 in BlaR1, eliminating the possibility for hydrogen bonding with Lys392 in a putative carboxylated state. Although the other available structures of class D β-lactamases show mutations at this position, they represent residues capable of hydrogen bonding [i.e. His73 in OXA-2 (PDB 1k38, unpublished), Ser73 in OXA-13 (Pernot et al., 2001)] or utilize a neighboring residue as a hydrogen bond donor [i.e. Ser120 in OXA-1 (Sun et al., 2003)]. Collectively, these differences in BlaRS create an environment that may discourage carboxylation of Lys392.  It is fascinating that these two distinctions are  conserved between BlaRS and class A β-lactamases such as SHV-2 (Figure 2.6c) as well as PBPs such as PBP2x (Figure 2.6d). In this sense, the active site of BlaRS best resembles a hybrid between that of the PBPs and the class D β-lactamases. 2.4.3  Formation of the Stable Penicilloyl Adduct BlaRS shares the greatest resemblance to the class D β-lactamases in terms of its fold  and sequence identity, but its sluggish deacylation activity is that of a PBP. Although carboxylation of Lys392 was not observed in the crystal structures of BlaRS, carboxylation of this residue under physiological conditions remains a cogent possibility.  As such, a  discussion of the mechanism of BlaRS must consider multiple scenarios, including those in which Lys392 is Nζ-carboxylated. In the observed case, where Lys392 is not carboxylated, the mechanism resembles the PBPs and requires an unprotonated Lys392 (Figure 2.7a) (Rhazi et al., 2003). Deprotonation of the lysine could be accomplished through the relatively hydrophobic environment of the Lys392 side chain consisting of Leu395, Met434, Phe442, Trp475, and Met476. Considering the hydrogen-bond network surrounding Lys392 in the apo-BlaRS 54  (a)  (b)  Figure 2.7:  Acylation schemes considering Lys392 as (a) non-carboxylated and (b)  carboxylated.  55  structure, including a tight hydrogen bond with the proposed nucleophile Ser389, a proton shuttling scheme similar to that proposed for the PBPs can be envisaged (Figure 7a). The pathway is initiated by positioning of the β-lactam antibiotic in the active site and abstraction of a proton from Ser389 by Lys392. The tetrahedral transition state (stabilized by the backbone nitrogens of Thr529 and Ser389) collapses to break the scissile β-lactam amide. The close proximity of Ser437 to two protonated lysines (Lys392 and Lys526) facilitates the abstraction of its hydroxyl hydrogen by the lone pair electrons of the β-lactam amide nitrogen.  Ser437 finishes the cycle by abstracting a proton from Lys392.  acylation, the Nζ of Lys392 is observed rotated away from the adduct.  Following  With Ser437  protonated and in the absence of a suitable general base for deacylation, BlaRS is stabilized in an acylated state. In an alternative mechanism, and as in the class D β-lactamases, the carboxylation of Lys392 in BlaR1 could provide a general base not only for acylation, but for deacylation as well. For this reason, a mechanism utilizing carboxy-Lys392 for acylation in BlaR1 must include a mode of preventing regeneration (i.e. deprotonation) of this residue as the general base for deacylation. Following a mechanistic scheme for the class D β-lactamases (Sun et al., 2003), deprotonation by the carboxylate of carboxy-Lys392 could activate Ser389 for nucleophilic attack of the β-lactam carbonyl (Figure 2.7b).  Were BlaRS an OXA-like  hydrolase, a deprotonated Ser437 could then subsequently deprotonate carboxy-Lys392 to regenerate its nucleophilicity and permit deacylation by activation of a bound water (Maveyraud et al., 2002). Without a structure showing a carboxylated Lys392 in BlaR1, it is difficult to rationalize how the position of the carboxy-lysine carboxylate would be perturbed in BlaR1 versus a class D β-lactamase. Indeed, what effect the lack of a stabilizing hydrogen 56  bond at the position of Leu395 and the substitution of an asparagine for valine at position 439 in BlaR1 can only be surmised.  Still, one possibility is that Asn439 may prevent  deprotonation of carboxy-Lys392 by accepting a hydrogen bond from the now protonated carboxylic acid group (Figure 2.7b). A molecule of water bound to Lys426 ultimately provides the proton for the shuttle instead, generating a stabilized acyl-enzyme. While Asn439 is not absolutely conserved in BlaR1/MecR1, this position appears to always be occupied by a polar residue capable of accepting a hydrogen bond [e.g. Thr452 in BlaR from B. licheniformis (Kerff et al., 2003)]. A third mechanistic possibility is that acylation encourages decarboxylation of Lys392 such that deacylation is prevented by removal of carboxy-Lys392 as a general base. This mechanism was first proposed to explain the acylated structures of BlaRS lacking carboxylation of Lys392 in addition to  13  C-NMR data showing BlaRS to have a diagnostic  carbamate peak (corresponding to carboxy-Lys392) that is lost upon acylation by ceftazidime (Birck et al., 2004). Decarboxylation of Lys392 upon BlaRS acylation was later confirmed by stopped-flow FT-IR kinetics and 13C isotope edited FT-IR spectroscopy (Thumanu et al., 2006). These experiments demonstrated for the first time that the acylation rate of BlaRS was severely attenuated in the absence of Lys392 Nζ-carboxylation. It was theorized that transfer of a proton from Ser389 to the Nζ of carboxy-Lys392 during acylation results in barrierless decarboxylation of Lys392, as supported by quantum-mechanical/molecular-mechanical (QM/MM) calculations (Birck et al., 2004). A similar branched scheme is proposed for OXA-10 based on QM/MM methods which is used to explain why supplemental NaHCO3 is required to simplify the biphasic kinetics of OXA-10 to monophasic at pH values below 7.5 (Li et al., 2005)—because NaHCO3 provides an excess of CO2 for facilitating  57  recarboxylation of Lys70 (Lys392 in BlaR1). Under this mechanism, deprotonation of Ser389 for acylation occurs via either Nζ or Oθ of carboxy-Lys392 (Figure 2.8). When Nζ of carboxy-Lys392 deprotonates Ser389, Lys392 is decarboxylated and BlaRS is trapped in its acyl-enzyme intermediate. When Oθ of carboxy-Lys392 deprotonates Ser389, carboxyLys392 is preserved as the general base for hydrolysis of the penicilloyl adduct. According to this mechanism, BlaRS should demonstrate some degree of residual β-lactamase activity and indeed BlaRS appears to hydrolyze 1 – 6 antibiotic molecules before being entrapped in the stable acyl-enzyme species observed in our crystal structure (Cha and Mobashery, 2007). In fact, the activity of BlaRS has been successfully altered from a PBP into a β-lactamase by conversion of Lys392 into S-(4-butanoate)-cysteine to produce an analog of carboxy-Lys392 which is unsusceptible to decarboxylation (Cha and Mobashery, 2007). Compared with the acylation mechanisms delineated in Figure 2.7, antibioticinduced decarboxylation appears to more comprehensively reconcile the available structural and functional observations. Indeed, the active site differences between OXA-10 and BlaRS previously described to putatively destabilize the carboxy-Lys392 carbamate are likely key to the function of BlaRS, rendering the acyl-enzyme irreversibly decarboxylated. Elucidating exactly how acylation contributes to lowering the stability of the carboxylated form of BlaRS would be greatly assisted by the structure of apo-BlaRS including a carboxylated Lys392, presumably acquired via crystallization at basic pH. 2.4.4 Structural Differences Between Apo- and Acyl-BlaRS The structures determined in this study provided us with the first opportunity to compare the two forms of the sensor domain of the BlaR1 receptor. The gross apo- and acylBlaRS structures are highly similar, possessing a rmsd in the range of 0.59-0.74 Å on the 58  Ser389  Ser389  O  O  O  H H  β-lactam  N  O  +  β-lactam  O  adduct  –CO2  NH2  (a) Lys392  Lys392  Ser389 O  X  H  H2O  O H  ..  N  –hydrolyzed antibiotic O  Ser389  Ser389 Lys392  (b)  O  O  β-lactam  OH H  N  O  Lys392  β-lactam  O  adduct  O H  N  O  Lys392  Figure 2.8: Acylation schemes considering protonation of Lys392 at (a) Nζ facilitating decarboxylation or (b) Oθ facilitating deacylation (modified from Cha and Mobashery, 2007).  59  231-239 Cα atoms. The four molecules that form the asymmetric unit of the apo-BlaRS structure are shown superimposed along with the two molecules that comprise the asymmetric unit of the acyl-BlaRS structure (Figure 2.9a).  It is clear from this  superimposition that the only regions that differ in position between the apo- and acyl-BlaRS structures are surface-exposed loops that likely vary due to thermal motion/mobility as opposed to significant conformational differences between the apo- and acyl-BlaRS forms. In the case of acyl-BlaRS, these loops exhibited weak electron density and higher-than-average B-factors.  These disordered regions include (i) the β-hairpin connecting strands 5 and 6  (residues 531-537), (ii) the large loop connecting α-helices C and E and including the short α-helix D (residues 406-427), and (iii) the N-terminal region up to β-strand 2 (residues 334360). While these loops were substantially better ordered in the apo structure, they made numerous stabilizing interactions with neighboring molecules in the P21 crystal lattice. The differences between the active sites of apo- and acyl-BlaRS are similarly subtle (Figure 2.9b).  Indeed, besides the actual acylation of Ser389, the largest structural  difference appears to be a slight repositioning of β-strand 5 due to hydrogen bonds with the benzylpenicillin adduct. The change in position of Ile531 is not surprising since it comprises the C-terminal end of β-strand 5 and the beginning of a hairpin turn that is poorly ordered in the acylated structure. A rotation about the χ1 of Thr527 is a consequence of a hydrogen bond with the carboxylate of the benzylpenicillin adduct. Acylation also reorients the Nζ of Lys392 away from Ser389, breaking the hydrogen bond observed in the apo structure.  60  (a)  (b)  Figure 2.9: Overlay of apo- and acyl-BlaRS. (a) Mainchain overlay represented as strings passing through Cα atoms. The blue strings correspond to the four molecules in the asymmetric unit of the apo-BlaRS structure while the green strings correspond to the two molecules in the asymmetric unit of the acyl-BlaRS structure.  The  benzylpenicillin adduct has been shown as yellow and purple sticks to indicate the variable position of the phenyl group. (b) Overlay of active site sidechains from the respective asymmetric units of apo- (tan) and acyl(green) BlaRS. The superimposition was generated with Swiss-Pdb Viewer “magic fit” (Guex and Peitsch, 1997).  2.4.5  Transmembrane Signal Transduction Circular dichroism (Golemi-Kotra et al., 2003) and FT-IR measurements (Thumanu  et al., 2006) have indicated an enhancement of secondary structure in S. aureus BlaRS upon acylation. In stark contrast, experiments with the BlaR sensor domain of B. licheniformis revealed no significant alterations in secondary or tertiary structure upon acylation with β-lactam antibiotics using a battery of biophysical techniques including circular dichroism, fluorescence spectroscopy, FT-IR, and deuterium/hydrogen exchange kinetics (Hanique et al., 2004). Considering these discrepancies between BlaR1 from S. aureus and BlaR from B. 61  licheniformis, it may be important to keep in mind that our structures were derived from S. aureus BlaR1. Our structures revealed no major conformational differences between the structures of apo- and acyl-BlaRS to provide an explanation as to how β-lactam-binding at the extracelullar face of BlaR1 initiates a signal capable of being transduced to the BlaR1 cytosolic protease domain. To date, there are now seven published sensor domain structures including three apo structures [B. licheniformis BlaR (Kerff et al., 2003), S. aureus BlaR1 (this work), MRSA MecR1(Marrero et al., 2006)] and four acylated structures (S. aureus BlaR1 + PenG (this work) / ceftazidime (Birck et al., 2004), MRSA MecR1 + PenG / oxacillin (Marrero et al., 2006)). These structures are all highly similar and reveal no significant conformational rearrangements upon acylation (Marrero et al., 2006). While we cannot rule out the possibility that the unexpected binding of pyrophosphate in the active site of our apo-BlaRS structure triggered the “acyl” conformation (indeed several of the interactions of pyrophosphate with the BlaRS active site closely resemble the interactions made with the benzylpenicillin adduct), if this were the case, distinct conformational differences should then be observed between these structures and the structure of the apo-BlaRS of B. licheniformis.  However, superpositions of the B.  licheniformis apo structure with the structures presented here reveal no significant conformational differences (1.23-1.27 Å rmsd on 214 Cα atoms for apo-BlaRS using pairwise comparisons with the molecules of the asymmetric unit and 1.07-1.22 Å rmsd for the same set of comparisons in acyl-BlaRS).  Similarly, two glycerol molecules were  apparently observed occupying the active site of the sensor domain of MecR1 without impairing the unbound conformation (Marrero et al., 2006).  While the presence of  pyrophosphate in the BlaRS active site is not expected to mimic acyl-BlaRS, it remains a  62  possibility that the acylated state is favoured without Nζ-carboxylation of Lys392. Even so, the subtle increase in secondary structure observed for BlaRS in solution upon acylation is not likely sufficient to fully account for signal transduction, especially considering the absence of this observation in the B. licheniformis BlaR sensor domain (Kerff et al., 2003). It is more probable that some component of the BlaR1 “sensor” remains missing. After BlaRS and the protease and transmembrane domains, the next largest domain in BlaR1 is the 56 residue extracellular loop (L2) connecting the second and third transmembrane helices. As the transmembrane domain is predicted to form a four-helix bundle, it is likely that L2 is in intimate contact with BlaRS and is thus excellently positioned to function as the missing trigger in signal transduction.  63  CHAPTER 3 – The BlaR1 L2 Loop & Signal Transduction  3.1  INTRODUCTION The crystal structures of the BlaR1 sensor domain (i.e. BlaRS) revealed no significant  conformational changes upon acylation of Ser389 by a β-lactam antibiotic (Chapter 2). In solution, however, circular dichroism and Fourier-transformed infrared spectroscopy (FT-IR) have implied that acylation of BlaRS produces an enhancement of secondary structure and reorientation into a more ordered, less dynamic state (Golemi-Kotra et al., 2003; Thumanu et al., 2006). Surprisingly these results contradict similar experiments and others performed using the sensor domain of BlaR from Bacillus licheniformis which indicated no such change in conformation (Hanique et al., 2004). Instead, the extracellular loop connecting the second and third transmembrane segments of BlaR (i.e. L2) was suggested to play a key role in transducing the acylation signal into the cytosol.  As evidence, a phage ELISA was  performed with the BlaR L2 loop displayed on the surface of a bacteriophage and the BlaR sensor domain attached to the ELISA plate. Retention of the L2-phage on the coated ELISA plate demonstrated a weak but specific interaction between the L2 loop and sensor domain of BlaR. Moreover, acylation of the BlaR sensor domain by washing the L2-bound ELISA plate with β-lactam antibiotics removed the L2-phage. Accordingly, Hanique et al. proposed that acylation of the BlaR sensor domain disrupts an interaction with the L2 loop, reorienting the transmembrane helices to activate the intracellular protease domain. This chapter describes several attempts to investigate the BlaR1 signal transduction mechanism. Firstly, NMR analysis was pursued to probe the conformational change that supposedly accompanies acylation of BlaRS by β-lactam antibiotics in solution. Mutation of 64  a conserved P-X-X-P motif in the L2 loop of BlaR1 was shown to prevent induction of β-lactamase expression in vivo, supporting the hypothesis that the L2 loop plays an important role in signal transduction. However, an interaction between BlaRS and the L2 loop could not be verified in vitro despite multiple attempts using constructs of the L2 loop prepared as a soluble domain.  3.2  METHODS  3.2.1  Expression and Purification of 15N and 2H/15N/13C-Labelled BlaRS 15  N-labeled and 2H/15N/13C-labelled BlaRS were produced starting with overnight  cultures of BL21 (DE3) E. coli transformed with pET41b(+)-BlaRS (Chapter 2) grown in LB medium containing 50 µg/mL kanamycin. For  15  N-labelled BlaRS, a 30 mL overnight  culture was centrifuged, washed with 15N-labelled M9 medium (prepared per L H2O with 6 g Na2HPO4, 3 g KH2PO4, 0.5 g NaCl, 120 mg MgSO4, 11 mg CaCl2, 2.7 mg FeCl3, 50 mg kanamycin, 1 mg thiamine, 10 g D-glucose, and 1 g 15  15  NH4Cl) and used to inoculate 2 L  N-labelled M9 medium for expression. The 15N-labelled culture was grown at 37 °C until  A600 ~ 0.6, heat-shocked at 42 °C for 1 h, cooled to 20 °C, and then induced to overexpress BlaRS overnight using 1 mM IPTG. For 2H/15N/13C-labeled BlaRS, a 50 mL overnight culture was centrifuged, washed with 2H15N13C-labeled M9 medium (prepared per L D2O with 6 g Na2HPO4, 3 g KH2PO4, 0.5 g NaCl, 120 mg MgSO4, 11 mg CaCl2, 2.7 mg FeCl3, 50 mg kanamycin, 1 mg thiamine, 3 g 13C-D-glucose, and 1 g 15NH4Cl) and used to inoculate 2 L 2H/15N/13C-labeled M9 medium for expression.  The 2H/15N/13C-labelled culture was  grown at 37 °C until A600 ~ 1.3 and then induced to overexpress BlaRS for 20 hrs at 37 °C using 1 mM IPTG. 65  The cells were lysed by high pressure homogenization using an Avestin EmulsiFlexC5. The  15  N-BlaRS protein was purified from the soluble cell fraction using three  chromatographic steps performed at 4 °C as described for unlabelled BlaRS in Chapter 2. 15  N-BlaRS in 20 mM Tris-HCl pH 7.5, 150 mM NaCl, 10% D2O was concentrated to 0.25  mM and stored at 4°C until analyzed. The 2H/15N/13C-BlaRS protein was purified from both the soluble and insoluble cell fractions to enrich yield. Insoluble material was resuspended in 6 M Guanidine-HCl, 40 mM NaHEPES pH 7.5, 0.2 M NaCl and dialyzed versus 40 mM NaHEPES pH 7.5, 0.2 M NaCl to refold 2H/15N/13C-BlaRS. Refolded and soluble cell lystate was then purified as described for unlabelled BlaRS (Chapter 2). The resultant protein consisted of pure 2H/15N/13C-BlaRS with a mix of amide 2H’s and 1H’s. To fully exchange the amide deuterons with protons, the pure 2H/15N/13C-BlaRS protein was first denatured by dialysis versus 6 M guanidine-HCl, 40 mM NaHEPES pH 7.5, 0.2 M NaCl and then refolded by a final dialysis versus 40 mM NaHEPES pH 7.5, 0.2 M NaCl. 2H15N13C-BlaRS in 20 mM Tris-HCl pH 7.5, 150 mM NaCl, 10% D2O was concentrated to 0.5 mM and stored at 4°C until analyzed. 3.2.2 NMR Data Collection and Assignment 1  H/15N-TROSY-HSQC spectra were acquired on  15  N-BlaRS in the absence and  presence of 1 mM PenG and 3D TROSY spectra (HNCA, HNCACB, HN(CA)CO, HNCO, HN(CO)CA, HN(CO)CACB) were recorded on 2H/15N/13C-BlaRS using Varian 500 Unity and 600 Inova spectrometers at 30°C (Kay, 2005; Sattler et al., 1999). NMR spectra were processed with NMRPipe (Delaglio et al., 1995) and analyzed/assigned with Sparky software (Goddard and Kneeler, 1999).  66  3.2.3  BlaR1 Mutant Constructs BlaR1 mutants were constructed by an overlapping PCR procedure as described  previously with the same end primers (Zhang et al., 2001). Mutagenic primers that were used in constructing BlaR1 C-terminal tags and proline point mutations are as follows: for His6 tagging,  P2h  (5’-CACCATCACCATCACCACGGTGTTTTAAATGGC-3’)  (5’-GTGGTGATGGTGATGGTGCATTTCTTTTAATATTTTTTCACTG-3’);  and  P3h  for  Myc  tagging, P2M (5’-GAACAAAAACTCATCTCAGAAGAGGATCTGAATTAGGGTGTTT TAAATGGCC-3’) and P3M (5’-CTAATTCAGATCCTCTTCTGAGATGAGTTTTTGTT CTTGGCCATTTAAAACACCC-3’); for mutation P49A, P2(P49A) (5’-GCAGGATTAATT GCTTTCATTCCTATT-3’) and P3(P49A) (5’-AATAGGAATGAAAGCAATTAATCC TGC-3’); and for mutation P52A, P2(P52A) (5’-GATTAATTCCTTTCATTGCTATTA AATTCTC-3’) and P3(P52A) (5’-GAGAATTTAATAGCAATGA AAGGAAT TAATC-3’). 3.2.4  β-Lactamase Assay A modification of the whole cell assay method of Kernodle et al. was used (Kernodle  et al., 1990). S. aureus cells were grown overnight at 37 °C in 2.5 mL of brain/heart infusion broth containing 10 µg/mL chloramphenicol to maintain the plasmid vector with and without 10 µg/mL CBAP as inducer on a reciprocal shaker platform at 250 rpm. 1 mL volumes were removed; cells were pelleted by centrifugation at 10,000 × g for 3 min; the broth was discarded; and cells were resuspended in 0.1 M sodium phosphate buffer (pH 6.0) to achieve a cell suspension of A570 = 0.95–1.05. 0.9 mL of bacterial suspension was added to 0.1 mL of 1 mM cephaloridine (final cephaloridine concentration of 100 µM) in sodium phosphate buffer and incubated at 37 °C. At 30 and 60 min, 1 mL samples were centrifuged to pellet cells.  The  cephaloridine  concentration  67  in  the  supernatant  was  determined  spectrophotometrically at 254 nm using buffer and the β-lactamase-negative control strain RN4220 as the blank. β-Lactamase activity was detected as a decrease in cephaloridine concentration. 3.2.5 Cloning and Purification of the L2 Loop The BlaR1 L2 loop (L2—amino acids 49-104; L2SOL—amino acids 54-104) was subcloned into a variety of expression vectors (Table 3.1 & Table 3.2). Standard cloning procedures were used except in the case of CBD-L2CYC-CBD where the Geiser et al. variation of the QuickChangeTM site-directed mutagenesis protocol (Geiser et al., 2001) was used to splice the L2 sequence into the pTWIN2 vector without the requirement for restriction enzymes. Expression conditions for the different L2 loop constructs varied in temperature and IPTG concentration. Typically, cells were grown from an overnight culture at 37 °C until A600 ~ 0.6 and then induced to overexpress 3-20 hrs at 15-37 °C using 0.5-1 mM IPTG. For L2, L2SOL and L2CH, the KSI-fusion proteins were expressed at 37 °C (1 mM IPTG) and purified using a 1 L scale up of the protocol provided in the pET Peptide Expression System 31 manual (Novagen). Briefly, the cells were suspended in 100 mL sonication buffer at pH 7.9 containing 5 mM imidazole, 40 mM Tris-HCl, 0.5 M NaCl, and two tablets of Complete EDTA-free protease inhibitors (Roche). Cells were then lysed by sonication on ice until no longer viscous and pelleted by centrifugation for 10 min at 12000 × g at 4 °C. The pellet was washed once by resuspension in 25-50 mL fresh sonication buffer and centrifugation. The insoluble pellet was then suspended in denaturing load buffer (pH 7.9) containing 6 M guanidine-HCl, 5 mM imidazole, 40 mM Tris-HCl, and 0.5 M NaCl and loaded at room temperature onto a pre-equilibrated 12-15 mL fast flow chelating sepharose  68  Table 3.1: Description of various L2 loop constructs. Construct  Sequence after purification*  Description  HL2  HHHHHHSSGLVPRGSHPFIPIKFSLF KFNNVNNQAPTVESKSHDLNHNINTT KPIQEFATDIHKFNWDSIDN  The native sequence of L2 from BlaR (Staphylococcus aureus) with a thrombin-cleavable N-terminal hexyl-His tag.  HL2 / BlaRS  HL2 (same as above)  HL2 co-expressed with BlaRS from a single plasmid to test the possibility that BlaRS may stabilize L2 expression  BlaRS (residues 330-585) KSI-L2  LPFIPIKFSLFKFNNVNNQAPTVESK SHDLNHNINTTKPIQEFATDIHKFNW DSIDN(HSe)  L2 expressed as a ketosteroid isomerase (KSI)-fusion to assist expression. The KSI domain is removed during purification leaving L2 with an N-terminal Leu and C-term homoserine (Hse).  KSI-L2SOL  LKFSLFKFNNVNNQAPTVESKSHDLN HNINTTKPIQEFATDIHKFNWDSIDN (HSe)  L2 expressed as a KSI-fusion but yielding a final product lacking the Nterminal “PFIPI” sequence that may reduce solubility.  KSI-L2CH  LCGGPFIPIKFSLFKFNNVNNQAPTV ESKSHDLNHNINTTKPIQEFATDIHK FNWDSIDNGGCLEHHHHHH  GFPL2SOL  MAHHHHHHSAALEVLFQGPGSKGEEL FTGVVPILVELDGDVNGHKFSVSGEG EGDATYGKLTLKFICTTGKLPVPWPT LVTTFSYGVQCFSRYPDHMKRHDFFK SAMPEGYVQERTISFKDDGNYKTRAE VKFEGDTLVNRIELKGIDFKEDGNIL GHKLEYNYNSHNVYITADKQKNGIKA NFKIRHNIEASKFSLFKFNNVNNQAP TVESKSHDLNHNINTTKPIQEFATDI HKFNWDSIDNNVDGSVQLADHYQQNT PIGDGPVLLPDNHYLSTQSALSKDPN EKRDHMVLLEFVTAAGITHGMDELYK  L2 expressed as a KSI-fusion but yielding a final product that includes Cys residues near the N- and C-termini to permit cyclization via an intramolecular disulfide bond. A C-terminal hexylhistidine tag was added to assist purification and interaction studies. With this construct, L2SOL was spliced into one of the soluble loops of GFPuv. Similar constructs have been made successfully for other peptides and even proteins. This fusion was intended to resemble L2 in its native state i.e. as the loop of a membrane protein. An N-terminal hexylhistidine tag was included to assist purification and interaction studies.  GFPL2  MAHHHHHHSAALEVLFQGPGSKGEEL FTGVVPILVELDGDVNGHKFSVSGEG EGDATYGKLTLKFICTTGKLPVPWPT LVTTFSYGVQCFSRYPDHMKRHDFFK SAMPEGYVQERTISFKDDGNYKTRAE VKFEGDTLVNRIELKGIDFKEDGNIL GHKLEYNYNSHNVYITADKQKNGIKA NFKIRHNIEASPFIPIKFSLFKFNNV NNQAPTVESKSHDLNHNINTTKPIQE FATDIHKFNWDSIDNNVDGSVQLADH YQQNTPIGDGPVLLPDNHYLSTQSAL SKDPNEKRDHMVLLEFVTAAGITHGM DELYK  69  GFP-L2SOL but with “PFIPI” reintroduced.  CBD-L2CYC-CBD  CRPFIPIKFSLFKFNNVNNQAPTVESKSHDL NHNINTTKPIQEFATDIHKFNWDSIDNA (N-term and C-term linked by peptide bond)  Another attempt to cyclize L2, this time by using the IMPACT-TWIN system (New England Biolabs).  (*) Amino acid residues not naturally in the L2 sequence are shown in bold. Table 3.2: Cloning of the L2 loop. Construct  p184R6H (10) HL2: pET28a-HL2  HL2  HL2 / BlaR  Source Plasmid  S  KSI-L2 KSI-L2SOL KSI-L2CH  S  BlaR : pET41bS BlaR pET28a-HL2 pET28a-HL2 pET28a-HL2  GFP: pGFPuv GFPL2SOL  L2SOL: pET28a-HL2  Primers  Target Plasmid  5’-GCATTACGCATATGCCTTTCATTCCTATT-3’ & 5’-GTAATGCCTCGAGTTAATTATCAATTGAATC-3’  pET28a(+) (Novagen)  His6L2: pET upstream primer #69214-3 (Novagen) & 5’-GTAATGCGGATCCTTAATTATCAATTGAATCCC-3’ S BlaR : ‘Cut and paste’ subcloning from existing restriction sites  pETDuet-1 (Novagen)  5’-GCATTACGCAGATGCTGCCTTTCATTCCTATT-3’ & 5’-CGTAATGCCTCGAGCATATTATCAATTGAATCCC-3’ 5’-GCATTACGCAGATGCTGAAATTCTCTCTTTTT-3’ & 5’-CGTAATGCCTCGAGCATATTATCAATTGAATCCC-3’ 5’-GCATTACGCAGATGCTGCCTTTCATTCCTATT-3’ & 5’-CGTAATGCCTCGAGGCAGCCACCATTATCAATTGAAT CCC-3’ GFP1-172: 5’-GCATTACGCCCGGGAGTAAAGGAGAAG-3’ & 5’-CGTAATGCGCTAGCTTCAATGTTGTGGCG-3’ L2SOL: 5’-GCATTACGGCTAGCAAATTCTCTCTTTTT-3’ & 5’-CGTAATGCAACGTTATTATCAATTGAATC-3’  pET31b(+) (Novagen) pET31b(+) (Novagen) pET31b(+) (Novagen)  CBDL2CYCCBD  NdeI/XhoI HL2: NcoI/BamHI S  BlaR : NdeI/XhoI AlwNI/XhoI AlwNI/XhoI AlwNI/XhoI  GFP1-172: XmaI/NheI pET47b(+) (Novagen)  L2SOL: NheI/AclI GFP173238: AclI/HindIII  GFP173-238: 5’-GCATTACGAACGTTGATGGATCCGTTCAAC-3’ & 5’-CGTAATGCAAGCTTTCATTATTTGTAGAG-3’ GFPL2  Restriction Sites  pET28a-HL2  5’-GCATTACGGCTAGCCCTTTCATTCCTATT-3’ & 5’-CGTAATGCACTAGTATTATCAATTGAATC-3’  pET47bGFPL2SOL (SpeI)*  NheI/SpeI  pET28a-HL2  5’-GTCGCGAATGACATCATTGTACACAACTGCCGCCCTTT CATTCCTATTAAATTCTC-3’ & 5’-CATTACAATGGTGTCACCGGATACGCATGCATTATCAA TTGAATCCCAATTAAAC-3’  pTWIN2  N/A**  (*) pET47b-GFPL2SOL was mutated to replace the non-unique AclI restriction site with unique SpeI to facilitate swapping the L2 sequence in place of L2SOL. QuickChangeTM site-directed mutagenesis was performed to replace 4 nucleotides using the following primers: 5’-GGGATTCAATTGATAATACTAGTGATGGATCCGTTCAACTAGC-3’ 5’-GCTAGTTGAACGGATCCATCACTAGTATTATCAATTGAATCCC-3’ (**) The above primers were used to create a PCR fragment spanning the L2 sequence with flanking regions matching the SapI cloning region of pTWIN2 and then inserted into pTWIN2 by QuickchangeTM mutagenesis without the requirement for restriction enzymes or DNA ligase (Geiser et al., 2001).  70  column charged with NiSO4. The column was washed for 5-10 column volumes in load buffer, then for 10 column volumes in wash buffer consisting of load buffer with 16 mM imidazole, then finally eluted in elute buffer composed of load buffer with 300 mM imidazole. Elution of the purified KSI-fusion protein was monitored by Coomassie stain and blue fractions dialyzed twice against 4 L H2O at 4 °C overnight to form a huge white precipitate. The precipitate was then pelleted by swing-bucket centrifugation at 3000 × g at 4 °C and the KSI fusion domain cleaved by CNBr digestion targeted at Met residues flanking the L2, L2SOL and L2CH sequences. To achieve digestion, the pellet was first dissolved by continuous stirring in 100 mL 80% formic acid either containing 1.0 g TCEP or bubbled with N2 (g) to protect Met residues from oxidation during CNBr digestion. 2.0 g of CNBr was then added and the reaction mixture stirred for 24-48 hrs in the dark. The reaction was stopped by rotovapping to dryness at 35 °C resulting in a clear gel. Whereas L2 was too insoluble to be efficiently purified from the cleaved KSI domain by selective solubilization, greater success was achieved with L2SOL using selective precipitation by dissolving the dry gel in load buffer and dialyzing vs. 1 L buffer containing 50 mM Tris-HCl pH 7.5, 0.5 M NaCl, and guanidine-HCl stepped down to 4 M, 3 M, 2 M, and finally 0 M. Insoluble and soluble  fractions  were  analyzed  by  tricine  polyacrylamide  gel  electrophoresis  (polyacrylamide: 4% stack, 10% spacer, 16% separation) and the final soluble product by MALDI-TOF mass spectrometry. L2CH was purified using selective solubilization followed by Ni2+-affinity. The dry gel was first dissolved in load buffer and then precipitated by dialysis vs. H2O. The precipitate was then selectively solubilized by stirring overnight at 4 °C in buffer at pH 7.5 containing 40 mM Tris-HCl, 0.2 M NaCl, 5 mM DTT, and 3 M urea. The soluble fraction was dialyzed overnight vs. 2 L load buffer and purified by Ni2+-affinity 71  as already described above. Pure L2CH was concentrated to ~4.5 mg/ml as estimated using a predicted 280 nm molar absorption coefficient of 5690 M–1 cm–1 (Pace et al., 1995). Concentrated L2CH was either used fresh (stored at 4 °C) or flash-frozen in liquid N2 and stored at -80 °C. For GFPL2SOL and GFPL2, expression was induced overnight at 28 °C using 1 mM IPTG. The cells were lysed in 100 mL load buffer (50 mM Tris-HCl, 20 mM imidazole, 0.3 M NaCl, pH 8.0) containing two dissolved tablets of Complete EDTA-free protease inhibitors (Roche) by high pressure homogenization using an Avestin EmulsiFlex-C5. GFPL2SOL was then purified by two chromatographic steps at 4 °C. The soluble cell lysate was then loaded onto a 15 mL pre-equilibrated Ni2+-chelated sepharose column, washed with ~10 column volumes of load buffer and eluted with elute buffer (50 mM Tris-HCl, 65 mM imidazole, 0.3 M NaCl, pH 8.0). The eluent was then passed through a Superdex 200 HR 10/30 size exclusion column (Amersham Biosciences) equilibrated in 20 mM Tris-HCl (pH 7.5) and 150 mM NaCl as the storage buffer. The dominant A280 peak was collected and concentrated to ~3 mg/ml as estimated using a measured 397 nm molar absorption coefficient of 38300 M–1 cm–1 (Pace et al., 1995). Concentrated GFPL2SOL was either used fresh (stored at 4 °C) or flash-frozen in liquid N2 and stored at -80 °C. For L2CYC, the L2 sequence is sandwiched between chitin-binding domains (CBDs) demarcated from L2 by inteins capable of self-cleavage. Expression of CBD-L2CYC-CBD was induced overnight at 20 °C using 0.5 mM IPTG. The cells were lysed in load buffer (20 mM HEPES, 0.5 M NaCl, 1 mM EDTA, pH 8.5) by passing the resuspended cells three times through a French pressure cell press. Following ultracentrifugation for 1 hr at 40000 × g to pellet insoluble cell debris, the soluble cell lysate was loaded onto 40 mL of pre72  equilibrated chitin resin (New England Biolabs) and washed with 5 column volumes of load buffer. The column was quickly flushed with 2-3 column volumes of load buffer adjusted to pH 7.0 and incubated overnight at room temperature to induce cleavage of the N-terminal CBD and liberation of an N-terminal Cys on the resin-bound L2CYC-CBD. The column was then washed with 2-3 column volumes of pH 7.0 load buffer and then quickly flushed with 23 column volumes of elute buffer (20 mM HEPES, 0.5 M NaCl, 1 mM EDTA, 50 mM 2mercaptoethane-sulfonic acid, pH 8.5) and incubated overnight at room temperature to cleave the C-terminal CBD and generate the C-terminal thioester for cyclization of L2CYC by intramolecular reaction with its N-terminal Cys. The next day, the column was eluted with just over a column volume of elute buffer and the eluent analyzed by tricine polyacrylamide gel electrophoresis. 3.2.6  Pull-Down and Crosslinking Assays with GFPL2SOL and BlaRS For the pull-down assay, 25 µL Nickel-charged chelating sepharose (SIGMA) was  equilibrated in binding buffer (50 mM Tris-HCl, 0.1 M NaCl, 20 mM imidazole, pH 8.0) and pelleted in a microcentrifuge at max speed. The resin was then incubated with 0.5 mL of 1 mg/mL His6-tagged GFPL2SOL in binding buffer for 10 min with agitation and washed three times with 0.5 mL aliquots of binding buffer. To bind BlaRS, GFPL2SOL-bound resin was incubated for 10 min with 0.5 mL of 1 mg/mL BlaRS in binding buffer and washed three times as previous. An additional wash step was then performed by incubating the resin in 2 mM PenG in binding buffer for 10 min or washed without PenG as a control. Following three additional washes with binding buffer, the resin was incubated in binding buffer containing 250 mM imidazole and 2 mM PenG or 250 mM imidazole alone. All steps were  73  performed at room temperature. Fractions of 5 µL were collected from the supernatant at appropriate steps during the assay and analyzed by 12% SDS-PAGE. For the cross-linking assay, the Lys-reactive cross-linker disuccinimidyl suberate (DSS) (Hill et al., 1979) was chosen to utilize the five Lys residues in the L2 sequence. Due to the high reactivity of DSS with amines, the Tris buffer in the samples was exchanged with a non-amine buffer (HEPES, 75 mM NaCl, pH 8.0). Prior to incubation with DSS, a mixture of BlaRS and GFPL2SOL was prepared at 0.5 mg/mL and incubated overnight at 4 °C. DSS stocks were prepared fresh in 50% DMSO immediately prior to setting up the assay. Crosslinking reactions were setup using 9 µL of protein and 1 µL of DSS and incubated for 10 min at room temperature. The reactions were stopped by the addition of 1 µL of quenching solution (1 M Tris, 1 M glycine, pH 7.5) and incubation at room temperature for 5 min. The samples were left on ice until analyzed by 12% SDS-PAGE.  3.3  RESULTS  3.3.1  NMR Spectral Assignments of BlaRS The conformation of BlaRS in solution was probed by NMR to investigate any  structural changes incurred by acylation at Ser389. The 1H/15N-TROSY-HSQC spectrum of apo-BlaRS differs substantially from the same spectrum following acylation of BlaRS with PenG (Figure 3.1) including altered chemical shifts for approximately 25% of the total 1  H/15N-TROSY-HSQC peaks. Considering the numerous interactions observed between  BlaRS and its PenG adduct in the crystal structure of acyl-BlaRS (Chapter 2), such differences could be explainable without a putative conformational change. Accordingly, we sought to assign the 1H/15N-TROSY-HSQC spectra of apo- and acyl-BlaRS in order to map 74  shifted peaks to the appropriate residues on the BlaRS crystal structure.  To facilitate  assigning the spectrum of a triply-labelled 30 kDa protein, TROSY-based triple-resonance  ω 1 - 15N  (ppm)  11  10  9  8  7  6  105  105  110  110  115  115  120  120  125  125  130  130  135  11  10  9  8  7  6  135  ω 2 - 1H (ppm) Figure 3.1:  Overlapped 1H/15N-TROSY-HSQC spectra showing the differences in chemical  shifts between apo-BlaRS (red) and BlaRS acylated by PenG (blue). An example of a shifted peak is magnified in the inset. The assigned peak list for apo-BlaRS is available in Appendix II.  experiments—with amide deuterons exchanged for protons—were utilized in assigning the signals from the main chain nuclei of apo-BlaRS. Unfortunately, only 45% of the 1H/15NTROSY-HSQC peaks for apo-BlaRS could be assigned unambiguously due to extensive signal broadening and spectral degeneracy. Many of the peaks that could be unambiguously assigned correspond to regions within the hydrophobic core of BlaRS, such as the large β-sheet of the α/β domain (Figure 3.2). Several of these assigned peaks were clearly shifted relative to their corresponding resonances in the 1H/15N-TROSY-HSQC spectrum of PenGacylated BlaRS and map to residues in close proximity to the PenG adduct (Figure 3.2). This  75  Assigned & Shifted Residues A396, L430, T432-V438, A455, W475-D478, T527-G528, G539-G543 Assigned & Unshifted Residues N359-N377, L395-I405, D429-V438, A455-G469, Y474-Q497, K506-N518, K526-G528, G539-T547, D550-G561  Figure 3.2: The crystal structure of PenG-acylated BlaRS demonstrating the proximity of the PenG adduct to residues corresponding to successfully assigned  1  H/15N-HSQC peaks.  Residues corresponding to assigned 1H/15N-HSQC peaks that were shifted upon acylation by PenG are highlighted in yellow, assigned but non-shifted in blue, and unassigned in grey. The PenG adduct is shown in magenta.  is hardly surprising considering that acylation of BlaRS by PenG would have a significant impact on the local electronic environment of active site residues and the residues immediately surrounding the β-lactam binding site.  However, these results are too  incomplete to dismiss allosteric changes in conformation upon acylation. Of course, even a complete map of all residues that were altered upon acylation would not reveal the structural nature of those changes. One possibility for characterizing the structural details of PenGacylation in solution is refinement of the acylated structure (obtained from crystallography) using residual dipolar couplings as restraints (Clore et al., 1998).  Unfortunately, this  technique would require a fully assigned 1H/15N-TROSY-HSQC spectra of apo- and acylBlaRS as a starting point. Alternatively, NMR relaxation experiments could potentially 76  provide insights into BlaR1 signal transduction by characterizing changes in the dynamics of apo and PenG-acylated BlaRS. 3.3.2 Constructs of the L2 Loop and Interaction with BlaRS The role of the L2 loop was probed in vivo in the S. aureus BlaR1 system by performing site-specific mutagenesis on highly conserved residues within the L2 loop. Mutation of either Pro49 or Pro52 to alanine in the L2 loop was observed to negate inducibility of β-lactamase expression (Table 3.3). In fact, β-lactamase activity was not even detectable in the proline mutants because cephaloridine concentrations were virtually identical to those of the β-lactamase-negative RN4220 control. Considering that BlaRS is alone sufficient to bind β-lactam antibiotics, the necessity of Pro49 and Pro52 for BlaR1 function suggests a critical role for the L2 loop in transducing the β-lactam-binding signal to Table 3.3: Summary of mutations and their effects on CBAP inducible β-lactamase expression assayed by hydrolysis of 100 µM cephaloridine. Mutation(s)*  30 min -I** (μM) +I (μM)  60 min -I (μM) +I (μM)  Phenotype  Controls RN4220 (negative) ZRI (wild-type)  100 ± 9 71 ± 6  98 ± 7 47 ± 14  100 ± 10 51 ± 10  102 ± 4 35 ± 7  Uninducible Inducible  C-terminal tags c[Myc] c[His6]  39 ± 4 45 ± 4  43 ± 7 46 ± 4  36 ± 3 38 ± 2  36 ± 1 40 ± 8  Constitutive Constitutive  102 ±2 100 ± 7  94 ± 9 99 ± 5  98 ± 5 99 ± 3  Uninducible Uninducible  Point Mutations in the L2 loop P49A P52A  98 ± 5 101 ± 9  (*) The S. aureus host strain RN4220 was transformed with a chloramphenicol-selectable S. aureus vector, pRN5542 (indicated as RN4220); with vector plus cloned wild-type blaR1-blaI-blaZ (ZRI); with vector plus wild-type blaI and blaZ and one of four blaR1 mutations, a C-terminal myc tag (c[Myc]), a C-terminal hexlyhistidine tag (c[His6]), an Ala-for-Pro substitution at residue 49 (P49A), and an Alafor-Pro substitution at residue 52 (P52A). (**) I indicates the presence of inducer, CBAP at 10 µg/ml.  77  the cell interior. Whether this role is directly related to signalling or a consequence of misfolding is unknown.  We have also observed that the fusion of either Myc or  hexylhistidine tags to the C-terminus of BlaR1 confers a phenotype of constitutive high-level β-lactamase expression (Table 3.3). To further characterize the function of the L2 loop in BlaR1, the L2 loop was expressed as a variety of deletion constructs (Table 3.1), spanning residues 49 to 104 and analyzed for expression, solubility, and potential interaction with BlaRS (Table 3.4). Multiple fusion constructs were tested to overcome the poor/absent expression commonly observed for polypeptides that are too short to contain sufficient secondary and tertiary structure to prevent proteolytic degradation in the cytosol. Expression as a fusion with the highly insoluble ketosteroid isomerase (KSI) domain stabilized the L2 loop by forcing it into inclusion bodies. Following removal of the KSI domain, the resulting L2 polypeptide could not be solubilized in an appropriate buffer so it was necessary to reclone the L2 loop lacking the hydrophobic N-terminus “PFIPI” to increase solubility (L2SOL). Since native L2 is the loop of a membrane protein and putatively has its termini held in close proximity by the fold of the BlaR1 transmembrane domain, cyclized L2 loop constructs may resemble native L2 more closely and thus bind more strongly to BlaRS in interaction assays. Indeed, considering the short length of the L2 loop (56 amino acids), proximal L2 termini may be essential to encourage formation of the L2 structural elements necessary for binding BlaRS. Several strategies were tested to cyclize the L2 loop including: (1) the addition of terminal Cys residues to permit a disulfide bond, (2) insertion of the L2 loop into a soluble loop of green fluorescent protein and (3) addition of an N-terminal Cys and C-terminal thioester to induce formation of a peptide bond between the L2 termini.  78  Table 3.4: Expression of various L2 loop constructs and when applicable the testing of an interaction with BlaRS.  Solubility  Interaction Test  Interact? (yes/no)  --  --  --  --  --  --  L2 (40%)  Insoluble  --  --  ~20mg/L Culture  L2SOL (>90%)  Low solubility  BlaRS+L2SOL native PAGE (EMSA)  no  ~20mg/L Culture  L2CH (>90%)  >4mg/mL  BlaRS+L2CH refolding and tested by gel filtration  no  (1) BlaRS+GFPL2SOL pull-down assay ±PenG  (1) no  (2) Cross-linking ±PenG  (2) no  Construct  Expression  HL2  None  HL2/BlaRS  poor, not reproducible  KSI-L2  ~20mg/L Culture  KSI-L2SOL KSI-L2CH  GFPL2SOL  ~1mg/L Culture  Product (purity) -HL2 not detectable after cell lysis  GFPL2SOL (>95%)  >3mg/mL  GFPL2  None  --  --  --  --  CBDL2CYC-CBD  Yes  no product after cleavage  --  --  --  Of the seven L2 constructs cloned, L2SOL (i.e. the L2-derived cleavage product of KSI-L2SOL), L2CH and GFPL2SOL were the only constructs successfully purified in a soluble form. Of these, L2CH and GFPL2SOL were the best candidates for interaction studies considering purity, solubility and the inclusion of a hexylhistidine tag for assisting with assaying a putative interaction with unlabelled BlaRS. GFPL2SOL had the additional advantage of ensuring that the L2 termini were proximally positioned (since green fluorescence was observed when GFPL2SOL was illuminated with ultraviolet light at 260 nm; Figure 3.3) whereas this was not easily verifiable for L2CH. Various assays were attempted with the available L2 constructs. Of these, L2SOL failed to produce a BlaRS band shift (i.e. the formation of a complex) in electromobility shift assays (EMSAs) and refolding L2CH with BlaRS failed to produce a stable complex via gel filtration under either reducing 79  or oxidizing conditions (to encourage disulfide bonding of the terminal Cys residues) (data not shown). Most convincingly, GFPL2SOL and BlaRS yielded only non-specific binding of BlaRS to Ni2+-sepharose/GFPL2SOL in pull-down assays (Figure 3.3) and no difference was observed between a cross-linked GFPL2SOL and BlaRS mixture and controls of these components cross-linked alone (Figure 3.4).  (i) (i) before expression  (a)  (ii) after GFPL2SOL expression  (ii)  (1) (2) (3) (4) (5) (6) (7)  (b)  97 66  (1) GFPL2SOL + BlaRS control  45  (2) GFPL2SOL flowthrough GFPL2SOL  31  S  BlaR  S (3) BlaR flowthrough  (4) Wash (5) 2mM PenG  21.5  (6) 250mM imidazole  15  (7) Imidazole + PenG  Figure 3.3: GFPL2SOL fluorescence and pull-down assay with BlaRS. (a) Fluorescence of E. coli cell pellets (i) before and (ii) after expression of GFPL2SOL (b) Pull-down assay using Nickel sepharose to bind His6-tagged GFPL2SOL. S  quantities of GFPL2SOL and BlaR used in the assay are shown in lane 1.  As a control, the Lanes 2 and 3  demonstrate binding saturation of the resin. Lanes 4 and 5 respectively show no difference between washing in the absence or presence of Penicillin G (PenG). difference between imidazole elution lacking or including PenG.  80  Likewise lanes 6 and 7 show no  (8) (9) (10) (11) (12) (13) (14) (8) No cross-linker  97 66  S (9) 1 DSS : 1 GFPL2SOL/BlaR mix  45  (10) 1 DSS : 1 mix, PenG GFPL2SOL  31  S  BlaR  (11) 10 DSS : 1 mix (12) 10 DSS : 1 mix, PenG (13) 50 DSS : 1 mix  21.5  (14) 50 DSS : 1 mix, PenG  15  (15) (16) (17) (18) (19) (20) (21) 97 66  (15) no cross-linker  45  (17) 1 DSS : 1 BlaR  (16) 1 DSS : 1 GFPL2SOL S  GFPL2SOL 31  S  BlaR  (18) 2 DSS : 1 mix (19) 2 DSS : 1 mix, PenG (20) 5 DSS : 1 mix  21.5  (21) 5 DSS : 1 mix, PenG  Figure 3.4: GFPL2SOL cross-linking assays with BlaRS. Cross-linking assays using the Lys-reactive cross-linker disuccinimidyl suberate (DSS).  Various  ratios of DSS were used in the presence/absence of PenG in an attempt to selectively trap the GFPL2SOL-BlaRS complex. Together, controls of GFPL2SOL and BlaRS cross-linked independently (lanes 16 and 17) combine to give the same pattern of bands as GFPL2SOL and BlaRS cross-linked as a mixture.  81  3.4  DISCUSSION  3.4.1  The Role of the L2 Loop in BlaR1 Signal Transduction The L2 loop has been shown to interact with the BlaR sensor domain from B.  licheniformis and has been implicated to play a critical role in β-lactam signal transduction (Hanique et al., 2004).  While it is generally assumed that the mechanism for signal  transduction in S. aureus BlaR1 will be highly similar to that of B. licheniformis, we have now demonstrated the essentiality of the L2 loop in the function of S. aureus BlaR1 by identification of two vital prolines which constitute part of a conserved P-X-X-P sequence motif located at the N-terminus of the L2 loop. Mutating either of these prolines severely attenuates β-lactamase expression, possibly by interfering with signal transduction. Since BlaRS is efficiently acylated by β-lactams even when expressed as a solitary domain (Golemi-Kotra et al., 2003), the requirement of the P-X-X-P motif in vivo suggests (as one possibility) that the L2 loop interacts with the sensor domain such that β-lactam-acylation of the sensor domain is “sensed” by the L2 loop as well. Accordingly, the nature of this interaction would determine basal levels of β-lactamase production, as was observed for the ZRI wild-type transformant which hydrolyzed some cephaloridine even in the absence of inducer. Since proline residues are fixed by an extra covalent bond to the protein main chain, these residues have less conformational freedom than any other amino acid. Considering this, the P-X-X-P motif would be inherently rigid and may serve to anchor the L2 loop appropriately for interaction with and regulation of the sensor domain. The lack of apparent conformational differences between the apo and acylated forms of BlaRS further supports that the L2 loop is responsible for modulation of BlaR1 activity by interaction with the BlaRS active site. However, it should be kept in mind that the membrane topology of BlaR1 (and 82  thus the positions of the L2 loop termini) is derived from studies in B. licheniformis, which predicted the P-X-X-P motif to lie at the N-terminus of the L2 loop (Hardt et al., 1997). Considering the hydrophobicity of the P-X-X-P motif, these residues are likely embedded at least partially with the lipid bilayer.  As such, these experiments do not rule out the  possibility that the P-X-X-P motif may be more involved in folding or membrane topology than signal transduction. While these results were not corroborated by our in vitro assays, the failure to observe an interaction between BlaRS and the L2 loop as non-contiguous constructs does not necessarily weaken the assertion that a critical interaction exists in BlaR1. Moreover, the low signal (15-fold higher than background) observed in the phage ELISA that identified the homologous interaction in B. licheniformis suggests that these domains interact relatively weakly (Hanique et al., 2004) and as such, observing this interaction may be highly sensitive to the experimental design.  83  CHAPTER 4 – The Modulation of MexR by ArmR  4.1  INTRODUCTION Bacterial multidrug efflux pumps confer widespread antibiotic resistance by actively  purging their cells of chemically diverse xenobiotics. The opportunistic human pathogen Pseudomonas aeruginosa possesses at least ten such efflux systems (Mima et al., 2007; Mima et al., 2005) which provide intrinsic resistance to numerous antimicrobials due to a synergy of heightened drug efflux and low outer membrane permeability (Nikaido, 1989; Poole et al., 1993; Poole and Srikumar, 2001). Based on their homology with the AcrABTolC efflux system in E. coli, drug efflux is likely accomplished via a rotating peristaltic pump-like mechanism powered by proton motive force (Murakami et al., 2006; Seeger et al., 2006). In P. aeruginosa, the MexAB-OprM efflux system displays the broadest substrate profile, including not only antibiotics (Li et al., 1995; Poole et al., 1993) and biocides (Chuanchuen et al., 2003), but organic solvents (Li and Poole, 1999; Li et al., 1998), dyes (Li et al., 2003), detergents (Srikumar et al., 1997) and homoserine lactones involved in quorum sensing (Evans et al., 1998; Pearson et al., 1999). Expression of mexAB-oprM is constitutive at basal levels in wild-type P. aeruginosa but mutations in any of three regulatory genes, mexR (Saito et al., 1999), nalC (Cao et al., 2004; Llanes et al., 2004; Srikumar et al., 2000) or nalD (Sobel et al., 2005), have been shown to produce mexAB-oprM hyperexpression and increased resistance to various medically relevant antimicrobials (Boutoille et al., 2004; Cavallo et al., 2002; El Amin et al., 2005; Pournaras et al., 2005). The primary regulator of the mexAB-oprM operon is the MarR family repressor MexR. MexR is a homodimer of 147 amino acids per subunit (total MW ~34 kDa) which 84  recognizes two operator sites that overlap promoters for both mexR and mexAB-oprM (Adewoye et al., 2002; Evans et al., 2001; Saito et al., 2001; Sanchez et al., 2002). The mexAB-oprM operon is additionally regulated at a weaker NalD-controlled promoter, but expression of mexAB-oprM is dominated by the promoter repressed by MexR (Morita et al., 2006). NalC indirectly influences mexAB-oprM expression by regulating a separate twogene operon, PA3720-PA3719 (Cao et al., 2004).  Upregulation of PA3719 alone (i.e.  without PA3720) has been shown to increase MexR protein levels without repressing MexAB-OprM production, suggesting that the small 53 residue protein encoded by PA3719 (MW ~6 kDa) can somehow alleviate the repressor activity of MexR (Cao et al., 2004). PA3719 appears to accomplish this via a direct protein-protein interaction with MexR (Figure 4.1) as demonstrated using a bacterial two-hybrid system and an in vitro transcription assay (Daigle et al., 2007). The gene encoding PA3719 has thus been renamed armR (anti-repressor for MexR). This distinguishes MexR as the first member of the MarR family shown to be modulated by a polypeptide effector as all other MarR family effectors known to date are small lipophilic compounds (Wilkinson and Grove, 2006). The MarR family of transcriptional regulators is widely distributed in bacteria and archaea and control a wide variety of biological processes, including resistance to antimicrobials, sensing aromatic compounds, and virulence (Ellison and Miller, 2006). While MarR proteins are poorly conserved in amino acid sequence, they share a common fold that consists of a helical dimerization domain and two winged helix (or winged helixturn-helix) DNA-binding domains (Alekshun et al., 2001). By conserving structure and evolving amino acid sequence, the different members of this family have apparently diverged  85  (1) ArmR is expressed due to an unknown signal(s) or NalC mutation  ?  ArmR  (3) β-Lactams & other molecules are extruded via MexAB-OprM efflux  NalC  OprM OM PA3720 armR  nalC  CW  MexAB-OprM MexR  MexA mexR  mexA mexB oprM  IM MexB  (2) ArmR alleviates repression of MexR, permitting expression of MexAB-OprM  H+  Figure 4.1: Schematic detailing regulation of MexAB-OprM-mediated efflux. Genes (shown as block arrows) and the proteins they encode are matched by color. Dotted arrows represent gene expression. 3D structures were derived from related published atomic coordinates (MexR – PDB ID 1LNW, MexA – PDB ID 1VF7, MexB – PDB ID 2J8S, OprM – PDB ID 1EK9).  to recognize a large variety of signaling molecules and DNA targets. The crystal structure of MexR (Lim et al., 2002) and a number of MarR proteins have been determined, but detailed mechanistic information is currently limited by a paucity of protein-ligand structures. This chapter investigates the role of ArmR as an allosteric effector of MexR. The binding of ArmR to MexR is shown to exclude DNA-binding and the MexR-binding region of ArmR is isolated to residues 41-53. To better understand how interaction with ArmR alleviates MexR repression, the crystal structure of the MexR double mutant Q106L/A110L  86  (MexRLL) was determined in complex with ArmR residues 29-53 (ArmRC). Comparison of this structure with the crystal structure of apo MexR reveals that ArmR induces a dramatic conformational change which repositions the MexR DNA-binding lobes into an orientation that is incompatible with binding DNA. This structure represents one of the first examples of a MarR family member bound by its effector and presents a compelling picture of an allosteric conformational change with important physiological consequences.  4.2  METHODS  4.2.1 Cloning and Purification of MexR and ArmR Wild-type MexR was produced by expression in BL21 (DE3) E. coli (Invitrogen, Carlsbad, CA) and purified via three chromatographic steps (SP-sepharose, heparin-agarose, superdex-75 gel filtration) as described previously (Lim et al., 2002). Purified MexR was stored in 20 mM Tris-HCl pH 7.5, 50 mM NaCl, concentrated to 50-100 mg/mL (as determined by Bradford assay), flash frozen in N2(l) and stored at -80°C until needed. The MexRLL-ArmRC complex was produced by co-expression of MexRLL with ArmRC followed by purification of the intact complex. The MexRLL expression plasmid, pET41a-MexRLL, was obtained through several rounds of QuickChangeTM site-directed mutagenesis, starting from the previously described wild-type MexR plasmid (Lim et al., 2002). These modifications produced a construct corresponding to MexR residues 1-142 and two point mutations, Q106L and A110L. The ArmRC expression plasmid, pTYB12-armRC, was constructed by subcloning ArmR residues 29-53 from plasmid PLC23 (Cao et al., 2004) into the BsmI-XhoI restriction sites of the IMPACT-CN vector pTYB12 (New England Biolabs, Ipswich, MA). BL21 StarTM (DE3) E. coli (Invitrogen), transformed with both 87  pET41a-mexRLL and pTYB12-armRC plasmids, was grown from an overnight culture at 37°C until A600 ~ 0.6, cooled to 20°C and then induced overnight using 0.1 mM isopropyl β-D-thiogalactopyranoside (IPTG). Purification of the complex was accomplished at 4°C via three chromatography steps: chitin affinity/intein tag cleavage, anion exchange and gel filtration. Cells were first resuspended in chitin load buffer (20 mM HEPES, 250 mM NaCl, 1 mM EDTA, pH 7.5) and lysed using a French pressure cell. The soluble cell fraction was loaded onto 50 mL pre-equilibrated chitin resin (New England Biolabs), washed with load buffer and then incubated overnight in elute buffer (20 mM HEPES, 50 mM NaCl, 1 mM EDTA, 50 mM DTT, pH 8.0) at 4°C to reduce the N-terminal tag of ArmRC to a single Ala residue. The next day, the column was eluted with fresh elute buffer and incubated for a second night to increase yield. The eluate was then diluted with buffer to reduce the NaCl concentration to 83 mM and applied to a Mono Q HR 10/10 anion exchange column (Amersham Biosciences, Piscataway, NJ). The complex was eluted over 100 mL with a linear 83–250 mM NaCl gradient in 20 mM HEPES, 1 mM EDTA, 14 mM β-mercaptoethanol, pH 8.0. Lastly, elution fractions containing protein were further purified via Superdex-75 gel filtration (GE Healthcare, Piscataway, NJ) in 20 mM Tris-HCl, 150 mM NaCl, 5 mM TCEP-HCl, pH 7.5. The pure complex was concentrated to 9.4 mg/mL using an Amicon Ultra-15 5K concentrator (Millipore, Billerica, MA) as estimated using a predicted 280 nm molar absorption coefficient of 12950 M–1 cm–1 (Pace et al., 1995), flash frozen in N2(l) and stored at -80°C until needed. ArmR and various truncated ArmR constructs were either expressed recombinantly or synthesized at >95% purity without using chemical ligation by C. S. Bio Company, Inc. (Menlo Park, CA) or the Brain Research Centre (Vancouver, BC, Canada). Recombinant 88  ArmR and ArmRC were separately cloned into the IMPACT-CN vector pTYB12 as described above. BL21 (DE3) cells carrying pTYB12-armR or pTYB12-armRC were grown from an overnight culture at 37°C until A600 ~ 0.5, cooled to 20°C, and then induced overnight using 0.1 mM isopropyl-β-D-thiogalactopyranoside (IPTG). The cells were lysed in load buffer (20 mM HEPES, 250 mM NaCl, 1 mM EDTA, pH 7.5) by passing the resuspended cells four times through a French pressure cell press. Following ultracentriguation for 1 hr at 40000 × g to pellet insoluble cell debris, the soluble cell lysate was loaded onto 25 mL of preequilibrated chitin resin (New England Biolabs) and washed with 15-20 column volumes of load buffer. The column was quickly flushed with 3 column volumes of elute buffer (20 mM HEPES, 50 mM NaCl, 1 mM EDTA, 50 mM DTT, pH 8.0) and incubated overnight at room temperature to induce cleavage of the N-terminal CBD. The next day, the column was eluted with just over a column volume of elute buffer. The eluate was further purified using a Mono S HR 5/5 cation exchange column (Amersham Biosciences) and a 0.05–1.0 M NaCl elution gradient in 20 mM HEPES, pH 8.0. Elution fractions corresponding to the principal A280 peak were combined and purified by semi-preparative C18 reversed phase HPLC using a 5–80% CH3CN gradient in 0.1% TFA. The major A280 peak was collected, lyophilized to dryness, and then dissolved in 20 mM Tris-HCl pH 7.5, 50 mM NaCl at a concentration of 0.3–2.0 M as estimated using a predicted 280 nm molar absorption coefficient of 9970 M–1 cm–1 (Pace et al., 1995). The identities of ArmR and ArmRC were confirmed by MALDITOF MS and frozen at -80°C. 4.2.2  Isothermal Titration Calorimetry (ITC) of MexR-ArmR ITC titrations were performed using a VP ITC (MicroCal Inc., Northampton, MA) in  20 mM HEPES pH 7.5, 50 mM NaCl by injecting consecutive 10 μL aliquots of 0.18 mM  89  ArmR (or truncated ArmR construct) into the ITC cell (volume=1.3528 mL) containing 29 µM MexR. Except for ArmRC, the ITC data were corrected for the heat of dilution of the titrant by subtracting mixing enthalpies for 10 μl injections of ArmR solution into buffer. For ArmRC the average heat of the last four data points was subtracted as background to correct for a slight difference in pH between the two solutions. The titration experiments were performed at 25°C to determine the binding constant of ArmR to MexR. Binding stoichiometry (N), binding enthalpy (ΔH), and equilibrium dissociation constants (Kd) were determined by fitting the corrected data to a single site interaction model (MicroCal Origin software). 4.2.3  Electromobility Shift Assay (EMSA) of MexR-ArmR EMSAs were carried out in 10 μL of 20 mM Tris-HCl pH 7.5, 50 mM NaCl as  described previously (Lim et al., 2002) using 0.6 nmol MexR in combination with 0.3 nmol double stranded operator DNA (DNAop) and/or an ArmR-dervied polypeptide at 0.6 or 1.2 nmol. Protein, DNAop, and polypeptide components were mixed and incubated overnight at 4°C prior to non-denaturing polyacrylamide (12% w/v) gel electrophoresis (PAGE) under reducing conditions at 80V for 2.5 hrs at room temperature. DNAop was prepared by mixing complementary oligonucleotides 5’-ATTTTAGTTGACCTTATCAACCTTGTTT-3’ and 5’-AAACAAGGTTGATAAGGTCAACTAAAAT-3’ (dissolved in 6×SSC buffer, 10 mM MgCl2, 1xTE buffer), denaturation at 96°C for 10 min and then cooling to 20°C at 0.5°C/min. Successful hybridization was confirmed by 20% PAGE in TBE buffer. 4.2.4  Limited Trypsinolysis of ArmR and MexR-ArmR Reactions were carried out on ice in 50 uL of cleavage buffer (20 mM HEPES pH  7.5, 50 mM KCl, 10 mM MgSO4).  MexR (2.9 nmol) or BSA (Sigma; 0.7 nmol i.e. 90  equivalent weight to MexR) was combined with synthetic ArmR (1.45 nmol) and the reaction started by addition of 0.5 µL of 5 mg/mL trypsin (Sigma). Reactions were stopped at various time points (t = 0 min to overnight) by transferring 1 µL to 9 µL of matrix solution (10 mg/mL alpha-cyano 4-hydroxycinnamic acid in 50% acetonitrile, 0.1% TFA) and analyzed using a Voyager DESTR matrix-assisted laser desorption ionization-time of flight (MALDITOF) mass spectrometer (Applied Biosystems) at the UBC MSL/LMB Proteomics Core Facility. Peptides were identified by comparison with a simulated tryptic digest (Baker and Clauser). 4.2.5  Expression and Purification of 15N/13C-Labelled ArmR 15  N/13C-Labelled ArmR was produced starting with a 60 mL overnight culture of  pTYB12-armR-transformed BL21 (DE3) E. coli grown in LB medium containing 100 µg/mL carbanicillin.  The overnight culture was centrifuged, washed with  15  N/13C-labelled M9  medium (prepared per L H2O with 6 g Na2HPO4, 3 g KH2PO4, 0.5 g NaCl, 120 mg MgSO4, 11 mg CaCl2, 2.7 mg FeCl3, 50 mg kanamycin, 1 mg thiamine, 3 g 15  13  C-D-glucose, and 1 g  15  NH4Cl) and used to inoculate 3 L  N/13C-labelled M9 medium for expression.  15  N/13C-labelled culture was grown at 37 °C until A600 ~ 0.5, cooled to 20 °C and then  induced to overexpress ArmR overnight using 0.1 mM IPTG. and lyophilized as described for unlabelled ArmR.  15  The  N13C-ArmR was purified  Dry doubly-labelled ArmR was  resuspended in 20 mM Tris-HCl pH 7.5, 50 mM NaCl, 10% D2O to a concentration of 1 mM, flash frozen in N2(l) and stored at -80°C until analyzed. 4.2.6 NMR Data Collection and Assignment 1  H/15N-HSQC spectra were acquired on 15N/13C-ArmR mixed with unlabelled MexR  and adjusted to pH 6.5 and 3D spectra (HN(CO)CACB, HNCACB, HN(CA)CO, HNCO)  91  were recorded on  15  N/13C-ArmR using Varian 500 Unity and 600 Inova spectrometers at  30°C (Kay, 2005; Sattler et al., 1999).  NMR spectra were processed with NMRPipe  (Delaglio et al., 1995) and analyzed/assigned with Sparky software (Goddard and Kneeler, 1999). 4.2.7  Crystallization and Structure Determination Crystals of the MexRLL-ArmRC complex were obtained by screening 480 microbatch  conditions at 8°C and 18°C using the Oryx 6 Crystallization Robot (Douglas Instruments Ltd., Hungerford, Berkshire, UK). The highest quality crystals grew as clusters from a mixture of 0.5 µL protein at 5 mg/mL and 0.5 µL 20% (wt/vol) polyethylene glycol (PEG) 3000, 0.1 M sodium citrate (pH 5.5) following two days under paraffin oil at 18°C. Single crystals were separated, cryoprotected in 20% (wt/vol) PEG 3000, 10% (wt/vol) PEG 1000, 0.1 M sodium citrate (pH 5.6) and flash frozen in N2(l). Data were collected to 1.8Å resolution at beamLine 8.2.2 of the Advance Light Source (Berkeley, CA) and processed with the program HKL2000 (Otwinowski and Minor, 1997). Despite the dramatic conformational differences between apo MexR and the MexRLLArmRC complex, the structure was successfully phased using molecular replacement (PDB ID 1LNW). To accomplish this, it was necessary to use only the dimerization domain of apo MexR (residues 3-31, 105-139) as an initial search model in the program Phaser (McCoy et al., 2007). The position of the dimerization domain was then fixed and the DNA-binding domain (residues 35-98) used as a search model in the program Molrep (Vagin, 1997). The structure was built using the program Coot (Emsley and Cowtan, 2004) and refined using the programs Phenix (Adams et al., 2002) and Refmac (Collaborative Computational Project, 1994) with a final R/Rfree of 17.6/22.9 (Table 4.1). Of the 310 residues in the complex, 129  92  residues were modeled in chain A of MexRLL (omitting disordered residues 63-65 and 8493), 135 residues were modeled in chain B of MexRLL (omitting disordered residues 62-67) and 24 residues were modeled in ArmRC (omitting N-terminal residue Arg29). Protein graphics were prepared with the program Pymol (DeLano, 2002) with the exception of Figures 4.7B and 4.9B, which were generated using Chimera (Pettersen et al., 2004).  4.3  RESULTS  4.3.1 MexR-ArmR Interaction Bacterial two-hybrid assays have indicated a direct interaction between MexR and ArmR in vivo (Daigle et al., 2007). To quantify this interaction in vitro, the binding of ArmR to MexR was analyzed by isothermal titration calorimetry (ITC) (Figure 4.2). Integration of the raw data (peaks of heat released upon binding of ArmR to MexR) gave an estimated stoichiometry of 0.45 moles of ArmR per mole of MexR, corresponding to one ArmR binding site per MexR dimer in solution, and a dissociation constant (Kd) of 290 nM. 4.3.2  ArmR Modulates MexR DNA Binding Affinity MexR regulates mexAB-oprM expression by binding to an operator region that  overlaps with the major promoter of this efflux operon (Evans et al., 2001; Morita et al., 2006). EMSAs involving a synthetic mexR operator sequence confirmed this MexR-DNA binding (Figure 4.3, lane 4). To function as an anti-repressor of MexR, ArmR must interfere with the ability of MexR to bind its cognate DNA operator. As expected from ITC, addition of ArmR to MexR yielded a MexR-ArmR complex (Figure 4.3, lane 5), while mixing ArmR, MexR and operator DNA produced complexes of both MexR-ArmR and MexR-DNA, but did not produce a ternary complex (Figure 4.3, lane 7). In fact, the MexR binding to  93  operator DNA appeared to be reduced slightly in the presence of ArmR (Figure 4.3, lane 6). The absence of a ternary complex consisting of MexR, ArmR and operator DNA demonstrates that MexR cannot simultaneously bind both ArmR and operator DNA, which is consistent with ArmR modulating MexR operator binding by sequestering MexR in complex with ArmR in lieu of a MexR-DNA interaction.  Kd = 290 ± 30 nM N = 0.45± 0.003 ΔH = -61 ± 0.60 kJ/mol ΔS = -80 J/mol·K  Figure 4.2: Binding of ArmR to MexR as measured by ITC. The upper panel shows the raw data of heat released by the binding of 10 µL injections of 0.18 mM ArmR into 29 µM MexR, while the lower panel shows the peak integration for each injection. The smooth red line displays the fit of the data to a single-site model. The equilibrium constant is reported for dissociation (Kd) and ΔH and ΔS correspond to the association of MexR and ArmR.  4.3.3  Isolation of the Minimal Peptide for Binding MexR Bacterial two-hybrid assays screening dozens of randomly-generated ArmR mutants  that failed to interact with MexR identified two mutants with single point mutations (Daigle et al., 2007). Interestingly, both of these mutations (L36P and W45A) were located within an 94  amphipathic C-terminal helix corresponding to residues 32-47 of ArmR as predicted by the JPRED server (Cuff et al., 1998) and helical wheel analysis.  An ArmR-derived sequence  spanning this region (residues 25-53) was confirmed to interact with MexR by two-hybrid assay (Daigle et al., 2007). Several strategies were pursued to further define the minimal ArmR sequence responsible for binding MexR.  ArmR DNA MexR  MexR-ArmR MexR MexR-DNA  MexR-DNA  DNA 1 Figure 4.3:  2  3  4  5  6  7  MexR-ArmR complex formation abrogates MexR-DNA binding.  EMSA  demonstrating the disruption of the MexR-operator DNA complex in lieu of formation of a MexR-ArmR complex as visualized by Coomassie brilliant blue staining of protein (top panel) and ethidium bromide staining of the MexR operator DNA (bottom panel). The ratios of MexR, operator DNA, and ArmR in each lane is indicated above the gel. (*) ArmR has a high pI and migrates in the opposite direction when not in complex with MexR. NMR spectral assignments of ArmR. Based on the prediction that only a subregion of ArmR binds MexR, we sought to further define this sequence by identifying peaks within the 1H/15N-HSQC spectrum of ArmR that shifted relative to their resonances in the 1H/15N-HSQC spectrum of the ArmR-MexR complex. The unassigned 1H/15N-HSQC 95  spectrum for free  15  N/13C-labelled ArmR (collected at pH 7.5) indicated a predominantly  random coil polypeptide with diagnostic amide chemical shifts in the range of 8-8.5 ppm (data not shown). The addition of MexR to this sample caused many peaks in the ArmR spectrum to disappear, presumably as a consequence of severe signal broadening due to the higher molecular weight of the MexR-ArmR complex.  The signal quality appeared to  improve after adjusting the pH to 6.5, but assignment of the resulting 1H/15N-HSQC spectrum only yielded peaks for a number of small fragments corresponding to the first 26 residues of ArmR. After purifying ArmR from the sample and analyzing it by MALDI-TOF mass spectrometry, it was deduced that the pH 6.5 spectrum corresponded to short N-terminal fragments of ArmR that were trimmed off the MexR-ArmR complex by microbial proteolysis. A single doubly-labelled peptide fragment corresponding to ArmR residues 25-53 (15N/13C-ArmR25-53) remained intact.  The identity of this peptide was  confirmed by assigning the 1H/15N-HSQC spectrum of this peptide, free of MexR (Figure 4.4). 15  Analysis of the secondary structure propensity (Marsh et al., 2006) of  N/13C-ArmR25-53 suggested a transient α-helix spanning residues 31-49 (Figure 4.5). As  with full-length ArmR, addition of unlabelled MexR to 15N/13C-ArmR25-53 resulted in a total disappearance of signal. Limited trypsinolysis of ArmR in complex with MexR. Trypsin cleaves peptide bonds C-terminal to arginine and lysine residues except in cases where these basic residues are immediately N-terminal to proline. The sequence of ArmR contains nine welldispersed arginine residues that are amenable to trypsinolysis, making trypsin ideally suited for assessing whether MexR preferentially protects a specific region of ArmR from limited proteolysis. Comparing the limited tryptic digests of ArmR in the presence of MexR versus  96  *  Figure 4.4: Assigned 1H/15N-HSQC spectrum of  15  N/13C-ArmR25-53. Observed peaks correspond  to residues 27-53 with residues 29-53 unambiguously assigned. Peaks for Ser25 and Ala26 are not observed as is typical for the first one or two N-terminal amino acids. The unassigned sidechains of Gln35 and Asn43 are shown connected by horizontal lines. Data was collected at pH 6.5, 30°C. (*) Aliased for 130 ppm.  97  Figure 4.5: Secondary Structure Propensity (SSP) of free  15  N/13C-ArmR25-53. A SSP score at a  given residue of 1 or -1 would reflect a fully formed α- or β-structure, respectively, while a score of 0.5 indicates that 50% of the conformers in the disordered state ensemble are helical at that position (Marsh et al., 2006).  BSA (as a control) suggests that the binding of MexR protects a C-terminal region of ArmR consisting of residues 31-53 and containing four arginines (Figure 4.6). A longer region spanning residues 25-53 was also somewhat protected. Arg12 appears to be protected even in the absence of MexR which suggests the presence of local structure in this region. ArmR truncations. Further characterization of the MexR-binding region of ArmR was pursued by measuring dissociation constants for MexR and a variety of truncated ArmR peptides (Figure 4.7). As indicated from limited trypsinolysis, truncation of 28 residues from the N-terminus of ArmR (ArmRC) had no effect on binding affinity. In fact, further N-terminal truncation up to Arg41 appeared to slightly improve binding affinity versus full-length ArmR.  In contrast, N-terminal truncation up to Tyr48 or C-terminal  truncation beginning at Tyr48 eliminated measurable binding, isolating the minimal MexRbinding region to within ArmR residues 41-53.  98  (A)  RDYTEQLR/DYTEQLRR  % Intensity  MSLNTPR  NKPSRTETEAVAASSGR  RNAWDLYGEHFY NAWDLYGEHFY SAVGRR DYTEQLR  MSLNTPRNKPSRTETEAVAASSGRSAVGRRDYTEQLRRAARRNAWDLYGEHFY  (B)  % Intensity  DYTEQLRRAARRNAWDLYGEHFY  RDYTEQLRRAARRNAWDLYGEHFY SAVGRRDYTEQLRRAARRNAWDLYGEHFY  Mass (m/z) Figure 4.6: MALDI-TOF mass spectra displaying limited tryptic digests of (A) ArmR with BSA and (B) ArmR with MexR. The full-length sequence of ArmR is shown between the two spectra with basic residues highlighted in large red type. The only lysine residue in the sequence is followed by a proline and therefore not an amenable site for trypsin cleavage. reactions on ice where no digestion was observed for MexR alone.  99  Spectra correspond to 1 min  Figure 4.7: Binding affinity of MexR with N- and C-terminally truncated ArmR constructs. All Kd values are from ITC experiments. An absence of an interaction is represented by a Kd >105 nM, which was the limit of detection for these experiments.  4.3.4  The Crystal Structure of the MexRLL-ArmRC Complex Initial attempts to co-crystallize the MexR with ArmR produced poor quality crystals  that were recalcitrant to standard optimization. Several strategies were pursued to improve diffraction quality, including truncating MexR and ArmR as well as engineering a hydrophobic crystallization epitope on the surface of MexR (Yamada et al., 2007). High diffracting crystals were finally obtained following deletion of five residues from the C-terminus of MexR and replacement of Gln106 and Ala110 with leucine (MexRLL) in addition to deletion of 28 residues from the N-terminus of ArmR (i.e. ArmRC). The rationale for modifying MexR was based on the structures of apo MexR which indicated these changes would not significantly alter the protein fold or function. Indeed, comparing the EMSAs of MexRLL vs. wild-type MexR reveals no noticeable differences in DNA or ArmR binding affinity (data not shown). The structure of the MexRLL-ArmRC complex was solved to 1.8 Å by molecular replacement and refined to a Rwork/Rfree of 17.6/22.9 (Table 4.1). Architecture of the MexRLL-ArmRC complex. The structure of MexR closely resembles other members of the MarR family and consists of two winged helix DNAbinding domains, each linked to a helical dimerization domain by a pair of long helices (Lim et al., 2002). As predicted by ITC, ArmRC binds the MexRLL dimer with a 1:1 stoichiometry 100  Table 4.1: Data collection and refinement statistics. Crystal Parametersa  Refinement Statisticsc  Spacegroup  P212121  Resolution Range, Å  48.56 – 1.80  Cell Dimensions: a × b × c, Å  52.2 × 57.2 × 91.9  Rwork / Rfree  17.6 / 22.9  Resolution, Å  1.8 (1.86 - 1.80)  rmsd bond lengths, Å  0.014  Wavelength, Å  1.00000  rmsd bond angles, °  1.401  No. Reflections  207187  No. Unique Reflections  26074  Redundancy  7.9 (8.0)  in parentheses  Completeness, %  100 (100)  b  Rsym = Σ|(Ihkl)-<I>| / Σ(Ihkl)  I/σI  35.1 (3.0)  c  5% of reflections excluded from  6.1 (5.6)  refinement  Rsym(%)  b  a  Highest resolution shell is shown  (Figure 4.8). ArmRC forms an α-helix spanning residues 32-49 with a severe 75° kink at Asn43.  ArmRC associates with MexRLL between the DNA-binding and dimerization  domains of MexRLL with residues C-terminal to the kink in the ArmRC helix (the C-terminal tail) buried between the two subunits of the MexRLL dimer.  Only one of the two  Leu106/Leu110 crystallization epitopes is involved in crystal contacts, forming a hydrophobic patch with His107 and symmetry-related Val126 and Ala129. As expected, these mutations are distant from ArmR and the DNA-binding domains. Interactions between ArmRC and MexRLL. The high affinity of MexR for ArmR is reflected in the crystal structure of the MexRLL-ArmRC complex, which reveals an extensive set of interactions focused around the C-terminal tail of ArmRC (Figure 4.9). These interactions are largely hydrophobic and include the bulky ArmRC sidechains of Trp45, Tyr48, Tyr53 and Phe52, which each fit into one of four hydrophobic pockets arranged in this order within the MexRLL interior. The first and last of these hydrophobic pockets (A and A’) are comprised of a nearly identical set of residues from subunits A and B 101  (A)  (B)  Figure 4.8: The crystal structure of MexRLL in complex with ArmRC. (A) Ribbon representation of the MexRLL-ArmRC complex (MexRLL in blue, ArmRC in orange). Disordered residues are shown as dashed lines. (B) Cross-section of MexRLL (blue surface) with ArmRC shown in ribbon and stick representation (C, N & O atoms in orange, blue & red, respectively).  102  Figure 4.9: Interactions between MexRLL and ArmRC. (A) Stick representation of ArmRC in its binding site (MexRLL C atoms in blue, ArmRC C atoms in orange; O, N & S atoms in red, dark blue & yellow, respectively). Hydrogen bonds are shown as dashed lines and ArmRC labels are italicized and colored orange to distinguish them from MexRLL. (B) Schematic map of interactions between MexRLL and residues 40-53 of ArmRC, showing hydrogen bonds and salt bridges by dashed lines. MexRLL residues involved in hydrophobic contacts are listed in boxes.  103  respectively, including Ile24, Leu28, Pro37, Val40 and Met112. The second and third hydrophobic pockets (B and B’) are made up of residues from both chains of MexRLL but likewise share a common set of residues, Phe17, Val20 and Met14. There are also a number of prominent polar interactions between ArmRC and MexRLL, including four hydrogen bonds to the backbone of ArmRC and eleven hydrogen bonds and salt-bridges to ArmRC sidechains. As with the hydrophobic interactions, identical polar residues from both chains of MexRLL are observed making similar interactions with residues in ArmRC. The sidechain amine of His116 (MexRLL) hydrogen bonds with the sidechain hydroxyl of Tyr48 (ArmRC) and the sidechain amine of His116’ (MexRLL) hydrogen bonds with the sidechain hydroxyl of Tyr53 (ArmRC). Additionally, the guanidinium group of Arg21 (MexRLL) makes an electrostatic interaction with the sidechain carboxylate of Asp46 (ArmRC) and the guanidinium group of Arg21’ (MexRLL) makes an electrostatic interaction with the backbone carboxylate of the ArmRC C-terminus. N-terminal to the kink at Asn43, the ArmRC helix is amphipathic and associates with the surface of MexRLL via a relatively hydrophobic interface. Prominent ArmRC residues in this interface include Thr33 which hydrogen bonds with Asp136 (MexRLL), Leu36 which contacts MexRLL residues Leu139 and Ala15’, as well as Ala40 which interacts with MexRLL residues Ala15’ and Met14’. The opposite face of the ArmRC helix is highly charged in this region due to the presence of six arginines, an aspartate and a glutamate.  4.4  DISCUSSION The high affinity of MexR for its effector ArmR (Kd = 290 nM) appears to be driven  by a large favorable heat release (ΔH = -61 KJ/mol), reflecting the large number of hydrogen  104  bonds, ionic interactions and van der Waals contacts observed in the MexRLL-ArmRC crystal structure. However, this net gain in enthalpy is opposed by a substantial (~1/3 as large) unfavorable loss in entropy (ΔS = -80 J/mol·K; -TΔS = 24 KJ/mol) which likely arises due to the induced folding of ArmR combined with the formation of a protein-peptide complex. Of course, these entropy losses would be offset by favorable entropy gains from the release of water caused by burying of the hydrophobic C-terminal tail. The affinity of MexR for its cognate DNA has not been measured, but other members of the MarR family demonstrate Kd values in the picomolar to nanomolar range (Wilkinson and Grove, 2006). This suggests that ArmR must express relatively strongly to fulfill its role as an anti-repressor. However, it should be noted that whereas the MexR-ArmR complex withstands size exclusion chromatography, MexR-DNA complexes appear to dissociate under identical conditions (data not shown), which indicates that MexR might prefer association with ArmR. 4.4.1 Structural Symmetry within ArmR Due to the symmetry of the MexR dimer, ArmR can bind MexR in one of two mutually exclusive orientations.  These sites cannot be occupied simultaneously as they  overlap in the interior region of MexR that binds the hydrophobic ArmR tail (Figure 4.10). This not only explains the stoichiometry of the MexR-ArmR complex, but also reveals a surprising capacity for structural symmetry in the C-terminal tail of ArmR despite an absence of symmetry in the amino acid sequence. Approximating the two orientations of ArmR by swapping the subunits of the MexRLL-ArmRC complex shows that the ArmR residues Trp45 and Phe52 occupy approximately equivalent positions, as do residues Tyr48 and Tyr53 (Figure 4.10). Consequently, the symmetry of the MexR dimer is largely preserved in the MexRLL-ArmRC complex despite extensive interactions with an asymmetric effector.  105  Indeed, the two MexRLL subunits superimposed with root mean squared deviations (rmsd) of 1.1 Å for all main chain atoms.  Figure 4.10: Overlapping binding sites reveal structural symmetry in the ArmR C-terminus. (A) Ribbon representation showing the position of a second ArmR-binding site (light orange) by swapping the A and B subunits of the MexRLL-ArmRC complex. (B) Hydrophobic pockets of the ArmRbinding site with MexRLL shown as a hydrophobic surface (green = hydrophobic) and residues 44-53 of ArmRC shown as (C, O & N atoms coloured orange, red & blue, respectively).  4.4.2 MexR and ArmR Mutants Several mutations have been reported to compromise the interaction between MexR and ArmR, including L35P, I104F, M112T, L135F, L28P and L75P in MexR as well as W45A and L36P in ArmR (Daigle et al., 2007). The importance of MexR residues Met112 and Leu28 are evident as these residues contribute to the hydrophobic pockets of the ArmR active site. Likewise, ArmR residues Trp45 and Leu36 are directly involved in MexR interactions. MexR residues Ile104, Leu35 and Leu75 are not directly involved in binding ArmR, but form a hydrophobic cluster in the hinge region between the DNA-binding domains and helices α1 and α5.  It is unsurprising that mutation of these residues to  dissimilar amino acids would interfere with binding ArmR as they are linked through van der Waal interactions with Leu28 to the ArmR A/A’ hydrophobic pockets. Leu135 is located in  106  the dimerization domain of MexR where it associates with its counterpart Leu135’ in the opposing subunit. Mutation of this residue to phenylalanine would thus introduce two bulky sidechains and potentially distort the dimerization interface. Considering the position of the ArmR-binding site between subunits of MexR, significant changes to the dimerization interface would be expected to impact the binding of ArmR. 4.4.3 ArmR-Induced Conformational Change The intrinsic conformational flexibility of MexR and other members of the MarR family is well established (Wilkinson and Grove, 2006). This flexibility is largely due to the plasticity of loops within the dimerization domain and the hydrophobic nature of the dimerization interface.  Moreover, while the winged helix DNA-binding domains are  relatively rigid entities, the flexibility of the dimerization domain results in various spacing between these two DNA-binding lobes. The crystal structure of apo MexR (Lim et al., 2002) provides an excellent example of this conformational flexibility, displaying four conformations of the MexR dimer with spacing between the two DNA-binding lobes ranging from 22.6-29.2 Å (Cα-Cα distance between Arg73 and Arg73’). An examination of each apo MexR conformation reveals that a conformational rearrangement is required to provide sufficient space for the bulky ArmR tail. This is clearly evidenced by overlapping the dimerization domains of MexRLL-ArmRC and apo MexR as severe steric clashes can be observed between ArmRC and the N-terminal residues of α2 and α2’ in apo MexR (Figure 4.11). To accommodate ArmRC-binding, each MexRLL subunit appears to have undergone a 20° helical bend in α5 that moved the C-terminus of this helix (i.e. Ala121) by 6-8 Å. In addition, α1 has pivoted at Thr22 to displace its N-terminus (i.e. Asp8) by 7-12 Å. These movements increase the distance between the dimerization and the  107  Figure 4.11: The binding of ArmRC induces a conformational change in MexRLL. (A) Comparison of the ArmR binding sites following superimposition of the dimerization domains of apo MexR dimer CD (yellow cylinders, PDB ID 1LNW) and MexRLL-ArmRC (blue cylinders with ArmRC shown as sticks coloured by atom type [C, orange; N, blue; oxygen, red; sulfur, yellow]).  (B)  Superimposition of the eight chains from the crystal structure of apo MexR (yellow ribbons) and the two subunits of MexRLL-ArmRC. (C) Superimposition of a single DNA-binding domain of apo MexR CD (yellow cylinders) and MexRLL-ArmRC (blue and orange cylinders).  108  DNA-binding domains of MexRLL and opens the A and A’ hydrophobic pockets. Concomitantly, ArmRC-bound MexRLL appears to have undergone a twisting of the DNAbinding domains with respect to the dimerization domain as well as each other. The outcome of this conformational change is a sheared orientation of the DNA-binding domains that shrinks the distance between the N-termini of the α4/α4’ recognition helices from 14.9-20.2 Å for apo MexR to 9.1 Å for the MexRLL-ArmRC complex (Cα-Cα distance between Leu67 and Leu67’). This displaces one domain relative to the other by 10.3-17.6 Å for the tip of the wing (i.e. Ser88) and 6.4-13.4 Å for the midpoint of the recognition helix (i.e. Arg73). 4.4.4 Allosteric Mechanism of MexR Anti-Repression Of the four conformations available for the apo MexR dimer, the widest spacing between the DNA-binding lobes was observed in dimer CD (Cα-Cα distance of 29.2 Å between Arg73 and Arg73’). Considering the spacing between major grooves in linear B-DNA is similar (34 Å), it was proposed that apo MexR dimer CD resembles the DNAbound conformation of MexR (Lim et al., 2002). Additionally, the shortest spacing was observed in dimer AB (Cα-Cα distance of 22.6 Å between Arg73 and Arg73’), which was found to bind the C-terminus of one of the other MexR molecules in the asymmetric unit. Based on these observations, it was speculated that a protein or peptide effector could modify the spacing of the DNA-binding domains and thereby regulate mexAB-oprM expression by dissociating MexR from its cognate DNA (Lim et al., 2002). It is now clear that this predicted effector binding site partially overlaps with the ArmR binding cleft.  The  sidechains of Asp146 and Ile147 from the MexR C-terminus appear to reasonably mimic the backbone carboxylate and sidechain of the ArmR C-terminus.  109  Currently, the only structure of a MarR family member in complex with its cognate DNA is OhrR from Bacillus subtilus [BsOhrR (Hong et al., 2005)]. The binding of BsOhrR to its pseudo-palindromic DNA operator was shown to induce a global bend of 10° and a slight under-twisting of the otherwise B-form DNA. These conformational changes appear to shorten the spacing requirement between the DNA-binding lobes of OhrR from 34 Å in linear B-DNA to 31.3 Å (Cα-Cα distance between Lys76 and Lys76’). Studies using atomic force microscopy with ExpG (Baumgarth et al., 2005) and circular dichroism spectroscopy with HucR (Wilkinson and Grove, 2005) suggest that similar protein-induced DNA conformational changes may be typical of the MarR family. This supports the proposal that apo MexR dimer CD—with a spacing between DNA-binding domains of 29.2 Å—may closely resemble its DNA-bound conformation. Indeed, a comparison of the structures for apo MexR dimer CD and DNA-bound BsOhrR (PDB ID 1Z9C) reveals that the orientation of the DNA-binding lobes is highly similar (rmsd of 2.2 Å for MexR Cα atoms corresponding to residues 37-99; Figure 4.12).  This conclusion is consistent with the  structures of reduced apo OhrR from Xanthomonas campestris [XcOhrR (Newberry et al., 2007)] and HucR (Bordelon et al., 2006), which have both been observed in conformations that are preconfigured for binding DNA. Using the apo MexR dimer CD as a model for the DNA-bound conformation, an allosteric mechanism for the anti-repression of MexR can now be proposed. The C-terminal tail of ArmR binds through a channel between MexR subunits and induces a sheared orientation of the DNA-binding lobes that is incompatible with binding DNA. Superimposition of one DNA-binding domain from apo MexR dimer CD with the MexRLL-ArmRC complex reveals a displacement of the α4’ recognition helix by 13.4 Å,  110  Figure 4.12: Mechanism of anti-repression. (A) Superimposition of the DNA-binding domains of apo-MexR dimer CD (PDB ID 1LNW; yellow cylinders) and DNA-bound bsOhrR (PDB ID 1Z9C; green cylinders with red ribbons) indicates a highly similar DNA-binding conformation for MexR.  (B)  Predicted wing movement and steric clashes with DNA using apo MexR dimer CD as a model for DNA-bound MexR superimposed with the DNA-binding domain of MexRLL-ArmRC.  resulting in severe steric clashes with the DNA backbone (Figure 4.12). Moreover, the DNA-binding wing is massively displaced from its position in the DNA minor groove (CαCα distance of 17.6 Å for Ser88). The wings of BsOhrR were observed making numerous minor groove interactions in the crystal structure of DNA-bound BsOhrR (Hong et al., 2005) and mutational analysis has established the importance of MexR wing residues Arg83 and Arg91 to DNA-binding (Saito et al., 2003). Finally, the position of ArmRC in its binding site projects the sidechain carboxylate of Glu50 into the gap between the two MexR DNA111  binding domains (Figure 4.11). This places a negatively charged moiety in close proximity to the helix-helix motif, a third DNA-binding element identified in the BsOhrR-DNA structure which is primarily composed of positively charged residues. As such, Glu50 may be involved in electrostatic repulsion of the negatively charged DNA backbone. 4.4.5 Comparison with Other MarR Family Members To date, MexR is the only MarR family member known to bind a protein effector. The ligands for the remaining MarR members are lipophilic (typically phenolic) compounds such as salicylate and uric acid (Wilkinson and Grove, 2006). In the case of OhrR, these lipophenolic molecules are organic hydroperoxides that induce structural changes by oxidizing a reactive cysteine. The recent crystal structure of oxidized XcOhrR revealed that OhrR proteins with multiple cysteines oxidize to form intersubunit disulfide bonds which stabilize a striking structural rearrangement of the DNA-binding lobes into a nearly perpendicular orientation (Newberry et al., 2007). This oxidized conformation appears to bear no resemblance to the sheared orientation of the MexRLL-ArmRC complex despite the nearby proximity of its reactive cysteine to several elements of the ArmR-binding site in MexR. The conformation of oxidized MarR members which utilize a single cysteine [e.g. BsOhrR and MgrA (Chen et al., 2006; Hong et al., 2005)] remains to be determined, but the orientation of XcOhrR appears unfavorable in the absence of a stabilizing intersubunit disulfide bond (Newberry et al., 2007). More likely, a conformational change is induced by formation of either cyclic sulfenamides or protein-effector mixed disulfides (Lee et al., 2007). The later possibility may induce rearrangements of the DNA-binding lobes via steric clashes with hydrophobic residues residing between the dimerization and DNA-binding domains, as observed with MexRLL-ArmRC.  112  The majority of MarR family members—such as MarR, HucR and EmrR—do not use reactive cysteines and are instead regulated by the non-covalent binding of low molecular weight ligands. In the first structure of a MarR family member, salicylate was observed bound to E. coli MarR in two sites (SAL-A and SAL-B) positioned on either side of the α4 recognition helix (Alekshun et al., 2001). This structure suggested a mode of regulation in which DNA-binding is prevented by occlusion of the DNA-protein interface.  The  physiological significance of either effector binding site could not be confirmed, however, as the SAL-A salicylate was involved in crystal contacts and the SAL-B salicylate was highly solvent exposed. Moreover, the broad conservation of these sites across the MarR family does not significantly support their importance in ligand-binding as SAL-A/B residues are likely involved in binding the DNA backbone, as revealed in the DNA-bound structure of BsOhrR. Interestingly, the conformation of MarR in the MarR-salicylate crystal structure closely matches that of MexRLL-ArmRC, superimposing to 2.8 Å rmsd (for 225 common Cα residues). This similarity includes not only the relative positions of the recognition helices, but also the large channel corresponding to the ArmR binding site. Despite a lack of conserved residues in this region, the presence of a channel indicates a potential effectorbinding site.  Indeed, recent structures of a MarR member from Methanobacterium  thermoautotrophicum, MTH313, revealed two unique binding sites for salicylate in this channel proximal to similar sites in XcOhrR (Saridakis et al., 2008). These binding sites appear to be mutually exclusive between the two subunits of the MTH313 dimer, producing an asymmetrical conformational change that enlarged the spacing of the DNA-binding lobes from 14 to 21 Å (unknown reference of measurement). From these data, it was posited that the apo form of MTH313 (with narrow spacing) represents the “active” conformation  113  (capable of DNA-binding) and the salicylate-bound structure (with wide spacing) represents the “inactive” conformation (incapable of DNA-binding). These conclusions are puzzling as they contradict the wide spacing observed in the structure DNA-bound BsOhrR. It is thus unfortunate that the coordinates for these structures are not presently available for comparison. While the coordinates for the MTH313 structures are presently unavailable for comparison, the salicylate-bound E.coli MarR structure is clearly ill-configured for binding a DNA double helix as its DNA-binding lobes share the same sheared orientation as the MexRLL-ArmRC complex. Regardless, the salicylate-binding sites identified in MTH313 appear to roughly correspond with the A/A’ hydrophobic pockets of MexR. This suggests that ArmR residues Trp45 and Phe52 may mimic the low molecular weight ligands of other MarR members and that Tyr48 and Tyr53 serve to anchor these moieties within the A/A’ pockets. It is intriguing from an evolutionary perspective that members of the MarR family characteristically interact with phenolic compounds as the sidechains of Tyr48 and Tyr53 (along with Trp45 and Phe52) occupy central positions in the hydrophobic ArmR binding site. Protein effectors offer several advantages over small molecule ligands, including the possibility to integrate multiple signals and increase binding specificity. As the only MarR protein currently known to be modulated a polypeptide effector, MexR is seemingly under tighter regulatory control than other members of the MarR family. Indeed, the affinity of MarR family members for their effectors typically produces Kd values in the micromolar range (Wilkinson and Grove, 2006). With a Kd of ~300 nM, the MexR-ArmR complex easily demonstrates the strongest such interaction in its class.  114  CHAPTER 5– Conclusions and Future Directions  The introduction of antibiotics revolutionized medicine, offering easy cures for previously untreatable diseases. However, resistance to each available antibiotic has been reported for every major infectious disease caused by bacteria (Avorn et al., 2001). We set out to understand in atomic detail how resistance to β-lactam antibiotics is controlled in disease-causing bacteria. Accordingly, two protein systems involved in regulating β-lactam resistance were investigated using a structural approach: the BlaR1 sensor/transducer from Staphylococcus aureus and the repressor MexR from Pseudomonas aeruginosa in complex with its polypeptide effector ArmR. Taken together, these studies touch on how each of the three modes of antibiotic resistance is regulated in bacteria. BlaR1/MecR1 controls the production of β-lactamase enzymes (i.e. drug destruction) as well as PBP2a (i.e. drug target modification) and MexR/ArmR regulates production of the MexAB-OprM multidrug efflux pump (i.e. drug expulsion).  5.1  THE BLAR1 SENSOR/TRANSDUCER FROM S. AUREUS The crystal structures of the S. aureus BlaR1 sensor domain (Chapter 2), including  apo (apo-BlaRS) and benzylpenicillin-acylated (acyl-BlaRS), constitute the first structures of the sensor domain of a BlaR1/MecR1 family member from S. aureus and the first sensor domain structure from any species exhibiting acylation with a β-lactam antibiotic. The previously published BlaR sensor domain structure from Bacillus licheniformis confirmed the predicted structural homology between BlaR1 family members and the class D β-lactamases (Kerff et al., 2003), but provided few mechanistic insights regarding β-lactam115  acylation or acylation-induced conformational changes and signaling due to the absence of an acylated structure. Even with the structures of apo- and acyl-BlaRS in hand, it was difficult to formulate a convincing mechanism whereby apo-BlaRS could be acylated by a β-lactam and then stabilized against deacylation as our structures showed an “active” arrangement of active site residues without Nζ-carboxylation of Lys392.  In fact, every subsequently  determined sensor domain structure has confirmed our active site configuration (Birck et al., 2004; Marrero et al., 2006). However, thanks to a wealth of supporting experiments (Birck et al., 2004; Cha and Mobashery, 2007; Golemi-Kotra et al., 2003; Thumanu et al., 2006), there is now compelling evidence to indicate that the BlaR1 family indeed utilizes carboxyLys392 for acylation and acylation-induced decarboxylation of Lys392 for trapping the acylenzyme intermediate in a covalent penicilloyl adduct. Elucidating exactly how acylation contributes to lowering the stability of the carboxylated form of BlaRS would be greatly assisted by the structure of apo-BlaRS including a carboxylated Lys392, presumably acquired via crystallization at basic pH. Our structures also provided the first opportunity to compare the apo and acylated forms of the sensor domain of a BlaR1 family member for conformational changes that might provide insights into the signal transduction mechanism of BlaR1. We observed no major movements of loops or other significant conformational changes induced by the acylation of Ser389, globally or proximal to Ser389. The conformation of the MecR1 sensor domain has now been shown to be similarly unaltered by acylation (Marrero et al., 2006). Multispanning transmembrane receptors such as BlaR1 are thought to relay their ligand-binding signals across the membrane via an allosteric twisting of a transmembrane helical bundle. In the case of BlaR1, this transmembrane domain is predicted to consist of a four-helix bundle  116  (Hardt et al., 1997) and acylation of BlaRS was proposed to break/alter a critical interaction between BlaRS and the L2 loop (Hanique et al., 2004), which somehow reorganizes the helices of the transmembrane bundle and its cytosolic loops to activate the zymogenic protease activity of the intracellular domain (Zhang et al., 2001).  By identifying two  essential prolines in the L2 loop, we have confirmed the L2 loop as a requirement for proper BlaR1 function in vivo (Chapter 3). We were unfortunately unsuccessful in our attempts to verify the proposed interaction between BlaRS and the L2 loop in vitro, but the lack of conformational differences between apo- and acyl-BlaRS underline the necessity for an additional component (such as the L2 loop) for transmembrane signal transduction. In the context of the entire BlaR1 signal transduction process, many questions remain that would clearly benefit from future structural studies, including structures of BlaRS in complex with the L2 loop, the intracellular protease domain both in its apo form and in complex with BlaI (or BlaR2), and ultimately full-length BlaR1 trapped in its various states. For the moment, structural investigation of full-length BlaR1 is paralyzed by an inability to acquire sufficient quantities of pure intact protein. S. aureus is among the most prevalent and antibiotic-resistant of pathogenic bacteria. Understanding the regulation machinery controlling β-lactamase and PBP2a expression in these Gram-positive bacteria may provide insights leading to the design of compounds that disrupt this regulatory pathway, restoring the activity of β-lactam antibiotics against drugresistant strains of staphylococci such as MRSA.  117  5.2  THE MEXR ANTI-REPRESSOR ARMR FROM P. AERUGINOSA The role of ArmR in P. aeruginosa as an anti-repressor of MexR has only recently  been proposed (Daigle et al., 2007) and is thus far completely unique among the MarR family of transciptional regulators. To function as an anti-repressor, ArmR must somehow neutralize the binding of MexR to its DNA operator. We have demonstrated for the first time that ArmR binds MexR with nanomolar affinity to the exclusion of operator DNA. To facilitate the crystallization of the MexR-ArmR complex for structural characterization, we attempted to isolate the minimal MexR-binding sequence in ArmR.  Part of this work  revealed that a 25-residue peptide derived from the C-terminus of ArmR (i.e. ArmRC) binds MexR just as tightly as full-length ArmR. The resulting crystal structure of MexRLL in complex with ArmRC provides the first structure of a MarR protein bound by a protein effector. The first observable residue in ArmRC (i.e. Arg30) is not involved in the ArmR helix (or any interaction) and is oriented away from MexR, indicating that residues Nterminal to it may likewise play little if any role in binding MexR. As such, the functional significance of these N-terminal residues (if any) remains to be determined. As a first step, examining the in vivo consequences in P. aeruginosa of deleting various N-terminal segments from ArmR may offer some insights into their role. By comparison with the previously determined structures of apo MexR (PDB ID 1LNW), the structure of the MexRLL-ArmRC complex reveals the allosteric conformational change responsible for alleviating repression of the mexAB-oprM operon. The structural symmetry inherent in the ArmR C-terminal tail induces a dramatic symmetrical rearrangement of the MexR DNAbinding domains which appears to render MexR incapable of binding DNA. Confirmation of this conclusion, however, will require a structure of MexR in complex with its cognate DNA  118  operator. Preliminary attempts not described here have indicated that accomplishing this will not be straightforward. There are presently few detailed structural descriptions of bacterial transcriptional regulators which utilize small protein effectors. Sporulation in Bacillus subtilis is regulated by the tetrameric repressor SinR and its 57 amino acid anti-repressor SinI. SinI forms heterodimers with SinR by mimicking residues in the hydrophobic interface between SinR subunits (Bai et al., 1993; Lewis et al., 1998). Carbon catabolite repression/regulation (CCR) in many Gram positive bacteria is controlled by the dimeric transciptional regulator CcpA (Henkin, 1996) and several allosteric effectors including phosphoproteins HPr (88 amino acids) and Crh (85 amino acids). Phosphorylated HPr or Crh act as corepressors and enhance the DNA-affinity of CcpA by binding as two monomers to opposite sides of the CcpA dimerization domain (Schumacher et al., 2004; Schumacher et al., 2006).  In addition,  carotenoid biosynthesis in Myxococcus xanthus is regulated by the dimeric repressor CarA and an 111 residue anti-repressor CarS (Lopez-Rubio et al., 2002; Whitworth and Hodgson, 2001). Unlike ArmR, CarS is highly acidic and thought to directly occlude the CarA DNAbinding domains in a 1:1 stoichiometry (Navarro-Aviles et al., 2007; Perez-Marin et al., 2004). In fact, the binding mode of ArmR appears to bare little resemblance to any peptide or protein effectors structurally characterized to date. Expression of armR is under the control of the NalC repressor, but the nature of the signal(s) to which NalC responds remains unknown. Considering that NalC belongs to the TetR family of transcriptional regulators (which all bind aromatic ligands), it is tempting to speculate that armR expression may be inducible by aromatic efflux substrates. In fact, while mexAB-oprM expression is generally considered to be constitutive, pentachlorophenol was  119  recently shown to upregulate both the PA3720-PA3719(armR) and mexAB-oprM operons (Muller et al., 2007) as well as a number of other efflux genes. Whether NalC responds directly to pentachlorophenol or to a secondary stress signal is currently unclear. In addition, the functional significance of the PA3720 gene that forms an operon with ArmR remains to be characterized as the deduced protein product of the PA3720 gene shows no significant sequence identity to any known protein and no observable phenotype is associated with either overexpression or loss of this gene (Cao et al., 2004). One possibility is that PA3720 plays a role in modulating NalC repressor activity in response to signals requiring expression of ArmR and, ultimately, mexAB-oprM. P. aeruginosa is a Gram-negative opportunistic pathogen that possesses expansive resistance to antibiotics due to the production β-lactamases and aminoglycoside-modifying enzymes, low outer membrane permeability, mutations in topoisomerases, and up-regulation of multidrug efflux pumps (Bonomo and Szabo, 2006). In fact, the accumulation of multiple resistance mechanisms in P. aeruginosa has spawned a number of “panresistant” strains with few susceptibilities (Deplano et al., 2005; Lolans et al., 2005). Deciphering the regulatory pathways controlling multidrug efflux pumps such as MexAB-OprM may provide clues about the physiological roles of these efflux systems, which appear to be involved in colonization and invasion of the bacterial host (Piddock, 2006). 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A protein modulator of multidrug efflux gene expression in Pseudomonas aeruginosa. J Bacteriol 189(15):5441-51.  Publications arising from B.Sc. studies 2004  Bateman, K. P., Baker, J., Wilke, M., Lee, J., Leriche, T., Seto, C., Day, S., Chauret, N., Ouellet, M. and Nicoll-Griffith, D. A. (2004) Detection of covalent adducts to cytochrome P450 3A4 using liquid chromatography mass spectrometry. Chem Res Toxicol 17(10):1356-61.  149  APPENDIX II – Assigned NMR Peak Lists for Apo-BlaRS  1  H/15N-TROSY-HSQC  Assignment ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN N359N-HN ?N-HN S360N-HN ?N-HN G361N-HN S362N-HN ?N-HN F363N-HN V364N-HN ?N-HN M365N-HN Y366N-HN S367N-HN ?N-HN M368N-HN K369N-HN K370N-HN D371N-HN K372N-HN  w1 106.074 106.666 107.018 107.382 110.541 112.309 113.452 114.621 114.563 114.541 114.567 114.651 114.544 116.017 116.536 117.113 117.112 115.938 115.213 116.954 117.822 117.333 119.744 114.92 116.019 115.271 111.194 111.883 115.457 120.115 127.67 117.612 124.644 126.295 120.891 119.579 126.511 121.716 115.105 121.104 117.225  w2 9.041 7.917 7.569 7.625 8.014 8.114 8.691 8.299 8.02 7.989 7.78 7.586 7.537 8.521 8.276 8.028 8.025 7.701 7.619 7.795 7.186 6.949 8.246 6.542 7.863 6.542 7.895 9.324 6.238 8.46 8.547 5.906 9.37 9.97 7.809 9.121 8.298 10.149 8.144 7.711 6.459  Assignment ?N-HN Y374N-HN I375N-HN ?N-HN Y376N-HN ?N-HN N377N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN L395N-HN ?N-HN A396N-HN M397N-HN F398N-HN ?N-HN G399N-HN ?N-HN L400N-HN ?N-HN D401N-HN ?N-HN R402N-HN H403N-HN I404N-HN I405N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN D429N-HN ?N-HN L430N-HN N431N-HN ?N-HN  150  w1 117.973 121.186 120.984 117.594 128.781 117.633 126.386 117.855 118.684 118.405 117.598 117.803 119.525 115.89 119.374 118.306 116.097 119.707 119.031 106.996 119.055 124.308 119.398 123.285 119.221 111.853 114.361 120.65 110.093 123.149 121.835 121.516 121.366 122.107 121.75 120.15 120.385 121.717 117.943 114.223 122.89  w2 8.726 9.799 9.054 8.619 9.942 8.429 8.933 8.158 7.788 7.765 7.65 7.574 7.695 6.585 7.815 7.748 6.815 8.71 7.372 8.809 7.31 8.565 7.284 9.147 6.988 7.92 7.682 8.576 6.902 9.512 9.097 9.1 8.992 8.847 8.64 8.547 8.009 8.438 8.053 7.522 8.51  Assignment ?N-HN A433N-HN M434N-HN ?N-HN Q435N-HN N436N-HN ?N-HN S437N-HN ?N-HN V438N-HN ?N-HN ?N-HN ?N-HN 442bN-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN ?N-HN A455N-HN ?N-HN T456N-HN Q457N-HN ?N-HN L458N-HN ?N-HN K459N-HN Q460N-HN L461N-HN ?N-HN N462N-HN ?N-HN Y463N-HN G464N-HN N465N-HN ?N-HN K466N-HN N467N-HN ?N-HN L468N-HN ?N-HN G469N-HN  w1  w2  119.432 126.017 117.477 121.859 117.019 112.381 120.705 117.526 122.861 119.044 119.778 119.958 119.921 119.735 120.92 120.871 122.959 123.312 122.726 121.689 120.21 122.623 120.104 116.477 120.538 121.306 116.801 121.318 118.847 120.531 116.674 122.95 116.018 122.358 118.911 113.062 120.12 123.023 112.878 117.006 124.571 125.079 121.909 106.246  8.899 8.614 8.288 8.357 8.169 7.745 8.333 7.581 8.248 8.493 8.224 8.182 8.071 8.096 8.138 8.072 8.023 8.047 7.873 7.86 7.887 8.767 7.786 7.608 8.071 7.737 8.94 7.73 7.607 8.152 8.626 7.466 7.645 7.183 8.29 6.689 8.723 6.979 8.375 7.187 5.949 8.45 5.671 7.959  Assignment T432N-HN ?N-HN ?N-HN Y474N-HN W475N-HN M476N-HN ?N-HN E477N-HN ?N-HN D478N-HN ?N-HN S479N-HN L480N-HN ?N-HN K481N-HN ?N-HN I482N-HN ?N-HN S483N-HN N484N-HN L485N-HN ?N-HN E486N-HN Q487N-HN V488N-HN I489N-HN ?N-HN V490N-HN ?N-HN F491N-HN K492N-HN N493N-HN ?N-HN M494N-HN ?N-HN M495N-HN ?N-HN E496N-HN ?N-HN Q497N-HN ?N-HN ?N-HN ?N-HN ?N-HN  151  w1  w2  114.033 131.963 129.444 115.459 120.846 119.864 128.082 121.424 125.865 122.13 127.852 122.201 126.219 128.381 122.355 126.747 120.906 126.49 123.98 123.342 117.004 124.553 114.296 116.685 116.661 119.095 126.754 116.932 125.848 120.504 118.065 118.313 123.44 120.629 124.126 112.541 123.406 116.06 124.489 116.509 125.046 124.835 127.172 130.171  8.664 10.324 9.59 6.297 6.631 6.579 9.131 8.05 9.064 8.293 8.997 9.153 7.29 8.826 8.358 8.681 9.828 8.654 7.828 9.853 8.966 8.635 7.367 8.36 6.9 7.443 8.44 9.145 8.346 7.145 7.875 8.179 8.342 8.499 8.119 7.941 8.076 8.636 8.013 7.61 7.988 7.891 7.941 7.851  Assignment ?N-HN K506N-HN N507N-HN Q508N-HN L509N-HN S510N-HN S511N-HN S512N-HN L513N-HN L514N-HN I515N-HN K516N-HN K517N-HN N518N-HN K526N-HN T527N-HN G528N-HN G539N-HN W540N-HN F541N-HN V542N-HN G543N-HN Y544N-HN V545N-HN I546N-HN T547N-HN D550N-HN K551N-HN Y552N-HN Y553N-HN F554N-HN A555N-HN T556N-HN H557N-HN L558N-HN S559N-HN D560N-HN G561N-HN ?N-HN ?N-HN ?N-HN ?N-HN  w1 117.978 120.656 117.547 120.268 118.436 116.48 114.905 117.82 120.253 121.399 129.348 115.756 121.335 122.845 118.143 116.68 106.289 111.263 119.962 118.74 127.119 110.808 114.871 118.884 126.715 119.517 119.334 123.517 122.897 122.053 118.946 119.832 122.137 125.052 123.914 113.756 119.175 111.306 122.707 116.678 115.045 118.831  w2 8.581 7.899 7.541 8.126 8.274 8.756 8.002 7.65 7.476 7.005 9.211 7.565 8.496 8.536 8.657 8.346 8.619 8.097 9.215 9.042 7.863 8.063 8.604 9.592 9.22 8.936 7.84 9.812 9.115 9.119 8.927 8.994 9.568 8.678 8.328 8.651 7.626 8.932 7.855 7.593 6.442 7.443  152  


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