New Antibiotic Targets (Methods in Molecular Medicine) [1 ed.] 9781588299154, 1588299155

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New Antibiotic Targets

M E T H O D S

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John M. Walker, SERIES EDITOR 142. New Antibiotic Targets, edited by W. Scott Champney, 2008 141. Clinical Bioinformatics, edited by Ronald J. A. Trent, 2008 140. Tissue Engineering, edited by Hansjörg Hauser and Martin Fussenegger, 2008 139. Vascular Biology Protocols, edited by Nair Sreejayan and Jun Ren, 2007 138. Allergy Methods and Protocols, edited by Meinir G. Jones and Penny Lympany, 2007 137. Microtubule Protocols, edited by Jun Zhou, 2007 136. Arthritis Research: Methods and Protocols, Vol. 2, edited by Andrew P. Cope, 2007 135. Arthritis Research: Methods and Protocols, Vol. 1, edited by Andrew P. Cope, 2007 134. Bone Marrow and Stem Cell Transplantation, edited by Meral Beksac, 2007 133. Cancer Radiotherapy, edited by Robert A. Huddart and Vedang Murthy, 2007 132. Single Cell Diagnostics: Methods and Protocols, edited by Alan Thornhill, 2007 131. Adenovirus Methods and Protocols, Second Edition, Vol. 2: Ad Proteins, RNA, Lifecycle, Host Interactions, and Phylogenetics, edited by William S. M. Wold and Ann E. Tollefson, 2007 130. Adenovirus Methods and Protocols, Second Edition, Vol. 1: Adenoviruses, Ad Vectors, Quantitation, and Animal Models, edited by William S. M. Wold and Ann E. Tollefson, 2007 129. Cardiovascular Disease: Methods and Protocols, Volume 2: Molecular Medicine, edited by Qing K. Wang, 2006 128. Cardiovascular Disease: Methods and Protocols, Volume 1: Genetics, edited by Qing K. Wang, 2006 127. DNA Vaccines: Methods and Protocols, Second Edition, edited by Mark W. Saltzman, Hong Shen, and Janet L. Brandsma, 2006 126. Congenital Heart Disease: Molecular Diagnostics, edited by Mary Kearns-Jonker, 2006 125. Myeloid Leukemia: Methods and Protocols, edited by Harry Iland, Mark Hertzberg, and Paula Marlton, 2006 124. Magnetic Resonance Imaging: Methods and Biologic Applications, edited by Pottumarthi V. Prasad, 2006 123. Marijuana and Cannabinoid Research: Methods and Protocols, edited by Emmanuel S. Onaivi, 2006 122. Placenta Research Methods and Protocols: Volume 2, edited by Michael J. Soares and Joan S. Hunt, 2006 121. Placenta Research Methods and Protocols: Volume 1, edited by Michael J. Soares and Joan S. Hunt, 2006

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M E T H O D S

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New Antibiotic Targets Edited by

W. Scott Champney Department of Biochemistry and Molecular Biology, East Tennessee State University, Johnson, Tennessee

© 2008 Humana Press Inc. 999 Riverview Drive, Suite 208 Totowa, New Jersey 07512 www.humanapress.com All rights reserved. No part of this book may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, microfilming, recording, or otherwise without written permission from the Publisher. Methods in Molecular MedicineTM is a trademark of The Humana Press Inc. All papers, comments, opinions, conclusions, or recommendations are those of the author(s), and do not necessarily reflect the views of the publisher. This publication is printed on acid-free paper.   ANSI Z39.48-1984 (American Standards Institute) Permanence of Paper for Printed Library Materials. Production Editor: Christina M. Thomas Cover design by: W. Scott Champney Cover illustration: The cover shows the chemical structures of a number of antibiotics currently used against some of the targets described in this book. Photocopy Authorization Policy: Authorization to photocopy items for internal or personal use, or the internal or personal use of specific clients, is granted by Humana Press Inc., provided that the base fee of US $30 per copy is paid directly to the Copyright Clearance Center at 222 Rosewood Drive, Danvers, MA 01923. For those organizations that have been granted a photocopy license from the CCC, a separate system of payment has been arranged and is acceptable to Humana Press Inc. The fee code for users of the Transactional Reporting Service is: [978-1-58829-915-4/08 $30 ]. Printed in the United States of America. 10 9 8 7 6 5 4 3 2 1 Library of Congress Cataloging in Publication Data: 2007933473

Preface A crisis is developing in the treatment of infectious diseases. Resistance of microorganisms to currently prescribed antibiotics is increasing at a steady rate, and fewer and fewer antimicrobial agents are available to treat these infections. Organisms displaying a multiple drug resistance phenotype are becoming common in both nosocomial and community-acquired infections. In some instances, there remains only a single antibiotic therapy suitable for treating the infection. This situation has stimulated several avenues of research in an effort to identify new drugs and new drug targets. Advances in combinatorial chemistry and high-throughput drug screening have provided a large number of compounds to be tested as new antimicrobial agents. The examination of microbial genome sequences has provided lists of potential new targets for such agents. This book is an attempt to bring these two areas together in the form of specific techniques that can be used to examine new drug targets and the effectiveness of new antibiotics. The major areas of vulnerability in microbial cells include cell wall and membrane synthesis and the processes of DNA replication, RNA transcription, and protein synthesis. Many of the currently used antibiotics have a specific inhibitory effect on one of these activities. Both new antimicrobial agents and modifications of existing drugs are being tested for improved antibacterial activity against these targets. In addition, a number of novel cellular targets are being examined as potential drug sites. These include the process of fatty acid biosynthesis, the activity of certain aminoacyl tRNA synthetases, the function of peptide deformylase, and the process of ribosomal subunit formation. Inhibitors of efflux pumps that lower cellular drug levels are also currently being sought. Additional potential drug targets include the inactivation of enzymes that modify antibiotics and the inhibition of enzymes that alter cellular antibiotic targets leading to resistance. Chapter 1 is a description of biocomputational methods for examining microbial genome sequences for novel inhibitory targets. Chapters 2 and 3 describe methodologies for examining inhibitors of DNA replication that affect topoisomerases (Chapter 2) and DNA polymerase III (Chapter 3). New assays for inhibitors of the bacterial RNA polymerase are discussed in Chapter 4. v

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Protein biosynthesis has been a major target for many different antimicrobial agents. Methods to target the aminoacyl tRNA synthetases are presented in Chapter 5. Chapters 6 and 7 present methods to examine inhibitors of bacterial ribosome biogenesis, a newly described target for antibiotics. Chapters 8 and 9 describe specific assays for inhibitors of ribosomal functions in translation including initiation (Chapter 8) and peptidyltransferase (Chapter 9). An additional promising new target is peptide deformylase; assays for inhibitors of its function are given in Chapter 10. The bacterial cell wall and membrane remain attractive targets for antibiotics. Chapter 11 discusses assays for penicillin-binding proteins of the cell wall, and Chapter 12 presents methods to assay inhibitors of lipopolysaccharide biosynthesis, an exciting new target. Membrane structure and function are the topics of Chapters 13 and 14, including measures of cationic antimicrobial peptide function (Chapter 13) and assays for changes in permeability using flow cytometry (Chapter 14). Chapter 15 describes several assays for detecting inhibitors of the ubiquitous efflux pumps in bacterial membranes. Two novel targets are described in Chapters 16 and 17. Inhibition of fatty acid synthesis (Chapter 16) is a promising new target, as are compounds that affect the important two-component signal transduction pathways in cells (Chapter 17). The remaining chapters describe assays for measuring target modification by ribosomal methyltransferases (Chapter 18), a discussion of inhibitors of -lactamase activity (Chapter 19), and assays for aminoglycoside-modifying enzymes (Chapter 20). The chapters in the book describe in detail specific methods that can be used to identify and assay these targets. As additional new compounds are identified and new targets in different microorganisms are found, these methods will aid in testing the antibacterial potential of these products. It is anticipated that this book will be primarily useful to laboratory investigators in academic, pharmaceutical, and medical institutions. W. Scott Champney

Contents

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Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Biocomputational Strategies for Microbial Drug Target Identification Kishore R. Sakharkar, Meena K. Sakharkar, and Vincent T. K. Chow . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Methods to Assay Inhibitors of DNA Gyrase and Topoisomerase IV Activities L. Mark Fisher and Xiao-Su Pan . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

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A Method to Assay Inhibitors of DNA Polymerase IIIC Activity Michelle M. Butler and George E. Wright . . . . . . . . . . . . . . . . . . . . . . . . . 25

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Methods to Identify and Characterize Inhibitors of Bacterial RNA Polymerase A. Simon Lynch and Qun Du . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 37

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Methods to Assay Inhibitors of tRNA Synthetase Activity Dieter Beyer, Hein Peter Kroll, and Heike Brötz-Oesterhelt . . . . . . . 53

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Three Methods to Assay Inhibitors of Ribosomal Subunit Assembly W. Scott Champney . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63

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Inhibition of Chaperone-Dependent Bacterial Ribosome Biogenesis Abdalla Al Refaii and Jean-Hervé Alix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75

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Assays for the Identification of Inhibitors Targeting Specific Translational Steps Letizia Brandi, John Dresios, and Claudio O. Gualerzi . . . . . . . . . . . . . 87

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SPARK: A New Peptidyl Transferase Activity Assay Alexander S. Mankin and Norbert Polacek . . . . . . . . . . . . . . . . . . . . . . . . 107

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High-Throughput Screening of Peptide Deformylase Inhibitors Kiet T. Nguyen and Dehua Pei . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117

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A Method to Assay Penicillin-Binding Proteins Michael J. Pucci and Thomas J. Dougherty . . . . . . . . . . . . . . . . . . . . . . . . 131

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A Method to Assay Inhibitors of Lipopolysaccharide Synthesis Marcy Hernick and Carol A. Fierke . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 143

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Methods for Assessing the Structure and Function of Cationic Antimicrobial Peptides Michelle Pate and Jack Blazyk . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155 14 Flow Cytometry of Bacterial Membrane Potential and Permeability Howard M. Shapiro . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 175 15

Bacterial Efflux Pump Inhibitors Barbara J. Kamicker, Michael T. Sweeney, Frank Kaczmarek, Fadia Dib-Hajj, Wenchi Shang, Kim Crimin, Joan Duignan, and Thomas D. Gootz . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187

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Mycobacterium tuberculosis -Ketoacyl Acyl Carrier Protein Synthase III (mtFabH) Assay: Principles and Method Sarbjot Sachdeva and Kevin A. Reynolds . . . . . . . . . . . . . . . . . . . . . . . . . . 17 Screening for Compounds That Affect the Interaction Between Bacterial Two-Component Single Transduction Response Regulator Protein and Cognate Promoter DNA Matthew G. Erickson, Andrew T. Ulijasz, and Bernard Weisblum . . 18 The Activity of rRNA Resistance Methyltransferases Assessed by MALDI Mass Spectrometry Stephen Douthwaite, Rikke Lind Jensen, and Finn Kirpekar . . . . . . . . 19 Assays for -Lactamase Activity and Inhibition Thammaiah Viswanatha, Laura Marrone, Valerie Goodfellow, and Gary I. Dmitrienko . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20 Studies of Enzymes That Cause Resistance to Aminoglycoside Antibiotics Engin H. Serpersu, Can Özen, and Edward Wright . . . . . . . . . . . . . . . .

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Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 273

Contributors Jean-Hervé Alix • Institute de Biologie Physico-Chimique, UPR 9073 du CNRS, University Paris 7 – Denis Diderot, Paris, France Abdulla Al Refaii • Institute de Biologie Physico-Chimique, UPR 9073 du CNRS, University Paris 7 – Denis Diderot, Paris, France Dieter Beyer • Bayer HealthCare AG, Wuppertal, Germany Jack Blazyk • Department of Biomedical Sciences, Ohio University College of Osteopathic Medicine, Athens, OH Letizia Brandi • Department of Biology MCA, University of Camerino, Camerino (MC), Italy Heike Brötz-Oesterhelt • AiCuris GmbH & Co. KG, Wuppertal, Germany Michelle M. Butler • Microbiotix Inc., Worcester, MA W. Scott Champney • Department of Biochemistry and Molecular Biology, East Tennessee State University, Johnson City, TN Vincent T. K. Chow • Human Genome Laboratory, Department of Microbiology, Yong Loo Lin School of Medicine, National University of Singapore, Singapore Kim Crimin • Pfizer Global R & D, MS, Groton, CT Fadia Dib-Hajj • Pfizer Global R & D, MS, Eastern Point Road, Groton, CT Gary I. Dmitrienko • Department of Chemistry, University of Waterloo, W. Waterloo, Ontario, Canada John Dresios • Department of Neurobiology, Scripps Research Institute, La Jolla, CA Thomas J. Dougherty • Pfizer Global Research and Development, Pfizer, Inc., Groton, CT Stephen Douthwaite • Department of Biochemistry and Molecular Biology, University of Southern Denmark, Campusvej, Denmark Qun Du • Cumbre Pharmaceuticals Inc., Dallas, TX Joan Duignan • Pfizer Global R & D, MS, Groton, CT Matthew G. Erickson • Department of Pharmacology, University of Wisconsin Medical School, Madison, WI Carol A. Fierke • Department of Chemistry, University of Michigan, Ann Arbor, MI L. Mark Fisher • Director, Molecular and Metabolic Signalling Centre, Division of Basic Medical Sciences, St George’s, University of London, Cranmer Terrace, London, UK Valerie Goodfellow • Department of Chemistry, University of Waterloo, W., Waterloo, Ontario, Canada Thomas D. Gootz • Pfizer Global R & D, MS, Groton, CT

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Claudio O. Gualerzi • Department of Biology MCA, University of Camerino, Camerino (MC), Italy Marcy Hernick • Department of Biochemistry, Virginia Tech, Blacksburg, VA Rikke Lind Jensen • Department of Biochemistry and Molecular Biology, University of Southern Denmark, Campusvej, Denmark Barbara J. Kamicker • Pfizer Global R & D, MS, Groton, CT Frank Kaczmarek • Pfizer Global R & D, MS, Groton, CT Finn Kirpekar • Department of Biochemistry and Molecular Biology, University of Southern Denmark, Campusvej, Denmark Hein Peter Kroll • Bayer HealthCare AG, Wuppertal, Germany A. Simon Lynch • Cumbre Pharmaceuticals Inc., Dallas, TX Alexander S. Mankin • Center for Pharmaceutical Biotechnology, University of Illinois, Chicago, IL Laura Marrone • Department of Chemistry, University of Waterloo, W., Waterloo, Ontario, Canada Kiet T. Nguyen • Wadsworth Center, New York State Department of Health, CMS, Albany, NY Can Özen • Graduate School of Genome Science and Technology, University of Tennessee, Knoxville, TN Michelle Pate • Department of Biomedical Sciences, Ohio University College of Osteopathic Medicine, Athens, OH Xiao-Su Pan • Department of Basic Medical Sciences, St George’s, University of London, Cranmer Terrace, London, UK Dehua Pei • Department of Chemistry and Ohio State Biochemistry Program, The Ohio State University, Columbus, OH Norbert Polacek • Innsbruck Biocenter, Division of Genomics and RNomics, Innsbruck Medical University, Innsbruck, Austria Michael J. Pucci • Achillion Pharmaceuticals, New Haven, CT Kevin A. Reynolds • Portland State University, Portland, OR Sarbjot Sachdeva • Portland State University, Portland, OR Kishore R. Sakharkar • Human Genome Laboratory, Department of Microbiology, Yong Loo Lin School of Medicine, National University of Singapore, Singapore Meena K. Sakharkar • Nanyang Centre for Supercomputing and Visualization, Nanyang Technological University, Singapore Engin H. Serpersu • Department of Biochemistry, Cellular and Molecular Biology, University of Tennessee, Knoxville, TN Howard M. Shapiro • The Center for Microbial Cytometry and Howard M. Shapiro, M.D., P.C., West Newton, MA Wenchi Shang • Pfizer Global R & D, MS, Eastern Point Road, Groton, CT Michael T. Sweeney • Pfizer Global R & D, MS, Eastern Point Road Groton, CT Andrew T. Ulijasz • Department of Pharmacology, University of Wisconsin Medical School, Madison, WI

Contributors

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Thammaiah Viswanatha • Department of Chemistry, University of Waterloo, W., Waterloo, Ontario, Canada Bernard Weisblum • Department of Pharmacology, University of Wisconsin Medical School, Madison, WI Edward Wright • Department of Biochemistry, Cellular and Molecular Biology, University of Tennessee, Knoxville, TN George E. Wright • GL Synthesis Inc., Worcester, MA

1 Biocomputational Strategies for Microbial Drug Target Identification Kishore R. Sakharkar, Meena K. Sakharkar, and Vincent T. K. Chow

Summary The complete genome sequences of about 300 bacteria (mostly pathogenic) have been determined, and many more such projects are currently in progress. The detection of bacterial genes that are non-homologous to human genes and are essential for the survival of the pathogen represent a promising means of identifying novel drug targets. We present a subtractive genomics approach for the identification of putative drug targets in microbial genomes and demonstrate its execution using Pseudomonas aeruginosa as an example. The resultant analyses are in good agreement with the results of systematic gene deletion experiments. This strategy enables rapid potential drug target identification, thereby greatly facilitating the search for new antibiotics. It should be recognized that there are limitations to this computational approach for drug target identification. Distant gene relationships may be missed since the alignment scores are likely to have low statistical significance. In conclusion, the results of such a strategy underscore the utility of large genomic databases for in silico systematic drug target identification in the post-genomic era.

Key Words: bacterial pathogens; comparative microbial genomics; essential genes; drug targets; antibiotics.

1. Introduction “Antibiotic” consists of the root words “anti,” meaning “opposed to” or “preventing,” and “biotic,” which is derived from the Greek word for life. In nature, various microbes and fungi secrete these compounds to gain an advantage in their microenvironment, and antibiotics are commonly isolated from such organisms. Between 1940 and 1960, the search for antibacterial agents relied From: Methods in Molecular Medicine, Vol. 142: New Antibiotic Targets Edited by: W. Scott Champney © Humana Press Inc., Totowa, NJ

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primarily on antibacterial activity, specifically growth inhibition of a desired spectrum of bacteria but not eukaryotic cells using in vitro laboratory assays. The molecular target was often identified only after the compound had reached the market. This was the “golden age of antibiotics,” during which the precursors of almost all the antibiotics that are in clinical application today were generated. Because antibiotics are such important medical compounds, the mechanisms by which antibiotics inhibit or kill bacteria have been under scrutiny for decades, with such studies being instrumental in the design of novel and improved compounds. The general modes of antibiotic activity are as follows: 1. Interference with the cell wall. Among the most well-known of antibiotics that interfere with peptidoglycan synthesis are the -lactams, exemplified by penicillin. 2. Interference with nucleic acid synthesis. For example, the sulfonamides (commonly known as sulfa drugs) mimic one of the folate precursors, competing with the precursor for the enzyme involved in the next step of folate synthesis, thereby hindering nucleic acid synthesis. Folate is an enzyme required for the synthesis of amino acids and nucleic acids. Other drugs that prevent nucleic acid synthesis are quinolones, which target bacterial DNA gyrase. Another class of drugs known as rifamycins specifically inhibits RNA synthesis by binding to RNA polymerase. 3. Interference with protein synthesis. These antibiotics inhibit the translational activity of the ribosome at various steps of protein synthesis. For example, puromycin is a non-selective inhibitor of protein synthesis that mimics tRNA. In addition, streptomycin causes the incorporation of incorrect amino acids at the “A” site of the growing polypeptide, whereas tetracyclines completely block protein translation by binding to a ribosomal subunit.

However, the widespread emergence of antibiotic resistance is of growing concern as an impediment in the therapy of bacterial diseases. In some instances, physicians are left with few or no antibiotics for the treatment of infections (e.g., Staphylococcus aureus), and such problems are likely to increase in magnitude. Clearly, a strategy of circumventing the threat of antibiotic resistance is to develop new drugs with novel targets and mechanisms of action. In the past decade, the tremendous progress in genome sequencing and analysis has complemented classical empirical approaches and has had a major impact on all biological sciences, including antibacterial research. Since the genome of Haemophilus influenzae was deciphered in 1995, many other bacterial genomes have been sequenced in rapid succession. These data pose a major challenge in the post-genomic era, i.e., to fully exploit this treasure trove for the identification and characterization of virulence factors in these pathogens, and to identify novel putative targets for therapeutic intervention (1). A critical advantage of this post-genomic analysis is the possibility of

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specifically identifying a target that is present in many or at least several bacterial genomes and/or one that may be susceptible to a “magic bullet” against a particular group of pathogens. Such bacterial genome analyses have revealed a variety of hitherto unexploited targets, with the potential for developing potent and selective antibiotics against a broad spectrum of bacterial pathogens. The target identification stage is the first step in the drug discovery process (Fig. 1) and as such can provide the foundation for years of dedicated research in the pharmaceutical industry. As with all the other steps in drug discovery, this stage is complicated by the fact that the identified drug target must satisfy a range of criteria to permit progression to the next stage. The possibilities of selecting targets through genomics-related methodologies are increasing, and several methodologies are currently available. An interesting approach designated “differential genome display” has been proposed for the prediction of potential drug targets (2,3). This approach relies on the fact that genomes of parasitic microorganisms are generally much smaller and encode fewer proteins than the genomes of free-living organisms. The genes that are present in the genome of a parasitic bacterium but absent in the genome of a closely related free-living bacterium are therefore likely to be important for pathogenicity and may be considered candidate drug targets. A complementary approach to target identification by bioinformatics has been reported in a concordance analysis of microbial genomes (4). A simple and efficient computational tool was developed (4) that can determine concordances of putative gene products showing sets of proteins conserved across one set of user-specified genomes but that are not present in another set of user-specified genomes (5). Recently, the use of fusion genes as putative microbial drug targets in Helicobacter pylori was suggested (6). Galperin and Koonin (7) proposed searching for drug targets

Fig. 1. Drug discovery pipeline.

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among previously characterized proteins that are specific and essential for a particular pathogen. Genes that are conserved in different genomes often turn out to be essential (8–11). A gene is considered to be essential if the cell cannot tolerate its inactivation by mutation, and its status is confirmed using conditional lethal mutants. Genomics can be applied to evaluate the suitability of potential targets using two criteria, i.e., “essentiality” and “selectivity” (4). The target must be essential for the growth, replication, viability, or survival of the microorganism, i.e., encoded by genes critical for pathogenic life stages. In order to address cytotoxicity issues, the microbial target for therapy should not have any well-conserved homolog in the host. This can help to avoid expensive dead ends when a lead target is identified and investigated in great detail only to discover at a later stage that all its inhibitors are invariably toxic to the host. Concurrently, the recent availability of the human genome sequence represents a major advance in drug discovery (12,13). Since this approach merely shortlists the putative microbial targets, the subsequent step would be to validate the prioritized targets, e.g., by constructing knockouts or by using chemical validation to verify their essentiality. Mutagenesis data are available for selected pathogenic microbial genomes and can be used to ascertain the essentiality of putative targets. Recently, Zhang et al. (14) compiled a list of all the currently available, experimentally determined essential genes into the Database of Essential Genes (DEG),

Fig. 2. Criteria for selection of proteins as drug targets.

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which includes the essential genes identified in the genomes of Escherichia coli, Haemophilus influenzae, Mycoplasma genitalium, Pseudomonas aeruginosa, Staphylococcus aureus, Vibrio cholerae, and Saccharomyces cerevisiae. The generation of a comprehensive gene list that is selective and essential to the pathogen will facilitate an accelerated genetic dissection of traits such as metabolic flexibility and inherent drug resistance in multiple pathogens (Fig. 2). Such a strategy will enable us to locate critical pathways and steps in pathogenesis; to target these steps by designing new drugs; and to inhibit the infectious agent of interest with novel antimicrobial agents. Here we discuss a novel in silico approach that provides a basis for addressing the “complexities and conundrums” in drug discovery by computational methods. 2. Materials 1. Microbial genome data and human genome data in relevant databases. 2. Computing power to perform comparative genome analyses (e.g., networked highend machine with large storage capacity). 3. Basic bioinformatics programs such as CD-HIT and BLASTP installed in the host machine.

3. Methods 1. Download the protein sequences for Homo sapiens and the bacterium of interest from the National Center for Biotechnology Information Website ftp://ftp.ncbi.nlm. nih.gov/genomes/ (15). 2. Download the Database of Essential Genes (DEG) from http://tubic.tju.edu.cn/ deg/ (15). 3. Download the CD-HIT program (16) from http://bioinformatics.ljcrf.edu/cd-hi. 4. Purge the bacterial genome file at 60% using CD-HIT to exclude paralogs from bacterium for further analyses. 5. Subject the resultant file to BLASTP against the H. sapiens genome to identify pathogen genes without homologs in humans (an expectation or E-value cutoff of 10−3 or lower is advisable). This will help to identify bacterial genes without human homologs and ensure that the target is selective for the genome of interest. 6. The non-homologous entries are then subjected to BLASTP against the DEG database for the identification of homologs to essential genes at a cutoff score of 10−10 . 7. The genes with hits are then classified into different groups based on gene names and metabolic pathways. Information on metabolic pathways can be obtained from the KEGG database available at http://www.genome.jp/kegg/.

Using the completely sequenced Pseudomonas aeruginosa genome as an example, the results demonstrate the unprecedented potential of the available

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complementary data sets (Table 1) and the application of a subtractive genomics approach (as explained above) for the identification of essential genes that may be considered as candidates for antibacterial drug discovery (4,17). Not surprisingly, many of the candidate genes code for the basic survival mechanisms of the bacterium (Fig. 3). The list of potential drug targets encoded in microbial genomes includes genes involved in translation, transcription, DNA replication, repair, outer membrane proteins, permeases, enzymes of intermediary metabolism, host-interaction factors, among many others. For example, the fatty acid synthesis pathway appears to be attractive for drug discovery given that a few known antibacterial compounds target enzymes of this pathway. Our strategy identified 11 genes from this group (18,19). Previous comparative analyses of complete genomes revealed that most of the pathogens have drastically diminished biosynthetic capabilities compared to their free-living relatives (20,21). Instead, these organisms depend on their hosts to provide essential nutrients such as amino acids, nucleotides, and vitamins. Thus, transport systems for these nutrients are generally well-conserved and easily identifiable (22). Analysis of metabolic pathways allows one to predict which substrates cannot be produced inside the cells and therefore need to be transported. This renders bacterial transport proteins that do not have human homologs as attractive drug targets (23). Our approach identified the category of “transport of small molecules” as a major one for drug target identification (9%). Protein translation, post-translational modification, and degradation were next on the list. Thus, these methods represent an efficient means for enriching potential target genes and for identifying those that are critical for normal cell functions (see Section 4).

Table 1 Results of the Computational Analyses of Pseudomonas aeruginosa Proteins Description Number of proteins Duplicates (>60% identical) Non-paralogs Number of proteins without hits in Homo sapiens Number of proteins with matches in DEG

Proteins 5567 119 5448 3841 306

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Fig. 3. Percentage distribution of 306 essential genes encoding different classes of proteins in Pseudomonas aeruginosa.

4. Notes It must be noted that this approach may not identify all the targets. Nonetheless, since gene disruption data are not available for all the genes in all the pathogens, this approach makes it possible to hazard a “first-order guess” for the probability that any untested gene is essential and may be a probable drug target. References 1 Miesel, L., Greene, J., and Black, T. A. (2003) Genetic strategies for antibacterial 1. drug discovery. Nat. Rev. Genet. 4, 442–456. 2 Huynen, M. A., Diaz-Lazcoz, Y., and Bork, P. (1997) Differential genome display. 2. Trends Genet. 13, 389–390. 3 Huynen, M., Dandekar, T., and Bork, P. (1998) Differential genome analysis 3. applied to the species-specific features of Helicobacter pylori. FEBS Lett. 426, 1–5.

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4 Sakharkar, K. R., Sakharkar, M. K., and Chow, V. T. (2004) A novel genomics 4. approach for the identification of drug targets in pathogens, with special reference to Pseudomonas aeruginosa. In Silico Biol. 4, 355–360. 5 Bruccoleri, R. E., Dougherty, T. J., and Davison, D. B. (1998) Concordance 5. analysis of microbial genomes. Nucleic Acids Res. 26, 4482–4486. 6 Sakharkar K. R., Sakharkar M. K., and Chow, V. T. (2006) Gene fusion 6. in Helicobacter pylori: Making the ends meet. Antonie van Leeuwenhoek 89, 169–180. 7 Galperin, M. Y., and Koonin, E. V. (1999) Searching for drug targets in microbial 7. genomes. Curr. Opin. Biotechnol. 10, 571–578. 8 Itaya, M. (1995) An estimation of minimal genome size required for life. FEBS 8. Lett. 362, 257–260. 9 Tatusov, R. L., Koonin, E. V., and Lipman, D. J. (1997) A genomic perspective 9. on protein families. Science 278, 631–637. 10 Koonin, E. V., Tatusov, R. L., and Galperin, M. Y. (1998) Beyond complete 10. genomes: From sequence to structure and function. Curr. Opin. Struct. Biol. 8, 355–363. 11 11. Kobayashi, K., Ehrlich, S. D., Albertini, A., Amati, G., Andersen, K. K., Arnaud, M., et al. (2003) Essential Bacillus subtilis genes. Proc. Natl. Acad. Sci. USA 100, 4678–4683. 12 Lander, E. S., Linton, L. M., Birren, B., Nusbaum, C., Zody, M. C., Baldwin, J., 12. et al. (2001) Initial sequencing and analysis of the human genome. Nature 409, 860–921. 13 Venter, J. C., Adams, M. D., Myers, E. W., Li, P. W., Mural, R. J., Sutton, G. G., 13. et al. (2001) The sequence of the human genome. Science 291, 1304–1351. 14 Zhang, R., Ou, H. Y., and Zhang, C. T. (2004) DEG: A database of essential 14. genes. Nucleic Acids Res. 32, D271–D272. 15 15. Wheeler, D. L., Church, D. M., Edgar, R., Federhen, S., Helmberg, W., Madden, T. L., et al. (2004) Database resources of the National Center for Biotechnology Information: Update. Nucleic Acids Res. 32, D35–D40. 16 Li, W., Jaroszewski, L., and Godzik, A. (2001) Clustering of highly homol16. ogous sequences to reduce the size of large protein databases. Bioinformatics 17, 282–283. 17 Stover, C. K., Pham, X. Q., Erwin, A. L., Mizoguchi, S. D., Warrener, P., 17. Hickey, M. J., et al. (2000) Complete genome sequence of Pseudomonas aeruginosa PA01, an opportunistic pathogen. Nature 406, 959–964. 18 Payne, D. J., Warren, P. V., Holmes, D. J., Ji, Y., and Lonsdale, J. T. (2001) 18. Bacterial fatty-acid biosynthesis: A genomics-driven target for antibacterial drug discovery. Drug Discov. Today 6, 537–544. 19 Heath, R. J., White, S. W., and Rock, C. O. (2002) Inhibitors of fatty acid synthesis 19. as antimicrobial chemotherapeutics. Appl. Microbiol. Biotech. 58, 695–703.

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20 Fraser, C. M., Gocayne, J. D., White, O., Adams, M. D., Clayton, R. A., 20. Fleischmann, R. D., et al. (1995) The minimal gene complement of Mycoplasma genitalium. Science 270, 397–403. 21 Lewis, K. (1999) Multidrug resistance: Versatile drug sensors of bacterial cells. 21. Curr. Biol. 9, R403–R407. 22 Clayton, R. A., White, O., Ketchum, K. A., and Venter, J. C. (1997) The first 22. genome from the third domain of life. Nature 387, 459–462. 23 Smith, R. S., Wolfgang, M. C., and Lory, S. (2004). An adenylate cyclase23. controlled signaling network regulates Pseudomonas aeruginosa virulence in a mouse model of acute pneumonia. Infect. Immun. 72, 1677–1684.

2 Methods to Assay Inhibitors of DNA Gyrase and Topoisomerase IV Activities L. Mark Fisher and Xiao-Su Pan

Summary DNA gyrase and DNA topoisomerase (topo) IV are the bacterial targets of coumarin and quinolone antimicrobial agents. Widespread resistance to clinically important antibiotics such as beta-lactams and macrolides has stimulated the development of novel gyrase and topo IV inhibitors especially against Streptococcus pneumoniae and other Gram-positive pathogens. Here, we describe how gyrase and topo IV activities are measured and how inhibitors of these enzymes may be assayed, focusing as a paradigm on DNA supercoiling by S. pneumoniae gyrase, DNA decatenation by S. pneumoniae topo IV, and DNA cleavage by both enzymes. These approaches provide mechanistic insight on inhibitor action and allow identification of dual gyrase/topo IV targeting agents that can minimize the emergence of bacterial resistance.

Key Words: DNA gyrase; DNA topoisomerase IV; DNA supercoiling; DNA decatenation; cleavage complex; ciprofloxacin; agarose gel electrophoresis; Streptococcus pneumoniae.

1. Introduction DNA gyrase and topo IV are important antimicrobial drug targets (1). Gyrase catalyzes the ATP-dependent supercoiling of DNA (2). The enzyme is present in all eubacteria but not in mammals and is unique among topoisomerases in promoting the negative supercoiling of DNA. Gyrase is essential for bacterial viability with functions in DNA replication, transcription, and recombination and in the regulation of chromosome supercoiling (1). First discovered in Escherichia coli, and best characterized from this source, gyrase has also been obtained in native or recombinant form from a variety of bacterial From: Methods in Molecular Medicine, Vol. 142: New Antibiotic Targets Edited by: W. Scott Champney © Humana Press Inc., Totowa, NJ

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species (3–5). Gyrase is a tetramer comprising two GyrA and two GyrB subunits encoded by gyrA and gyrB genes, respectively. Its mechanism of DNA supercoiling proceeds via the passage of one DNA segment (the “transported” or T segment) through a transient double-stranded break in a second DNA helical region (the “gate” or G segment) involving a covalent enzyme-DNA complex called the “cleavage complex” (6–8). Strand passage is achieved through the concerted opening and closure of protein–protein and protein–DNA gates. It is believed that the T and G segments lie within a 120–150-bp region of DNA wrapped on the enzyme such that the T segment is presented in the correct orientation to generate negative supercoiling on strand passage (6–8). Gyrase shares homology and mechanistic similarity with topo IV (9), a ParC2 ParE2 tetramer that also acts via a double-stranded DNA break to facilitate ATP-dependent chromosome decatenation and relaxation. Both enzymes change DNA linking number in steps of two and are categorized as type II topoisomerases (6–8). In addition to its biological and mechanistic interest, gyrase is the target for coumarins, quinolones, and other novel classes of antimicrobials (10–13). Coumarins, e.g., novobiocin, as well as the cyclothialidines and aminobenzimidazoles, inhibit DNA supercoiling by acting as competitive inhibitors of the GyrB ATPase (10,11,13). By contrast, quinolones and their fluoroquinolone analogues such as ciprofloxacin inhibit gyrase activity by stabilizing the cleavage complex involving the GyrA subunits (1,12). The traditional view of gyrase as the primary quinolone target holds for Gram-negative bacteria such as E. coli but not for Gram-positive species such as S. pneumoniae, wherein the target can be gyrase or topo IV, or both, depending on the drug structure (14,15); see also (13). Ongoing development of novel agents (13) and the recent clinical introduction of new quinolones such as levofloxacin, moxifloxacin, gatifloxacin, and gemifloxacin directed specifically against Gram-positive pathogens (16) have identified the need to understand how inhibitors interfere with gyrase and topo IV. The assay for gyrase activity involves monitoring ATP-dependent supercoiling of a relaxed circular plasmid DNA (3,4). The reaction products are separated and displayed by agarose gel electrophoresis. Supercoiled DNA is more compact than relaxed DNA and therefore migrates more rapidly through the gel. Though almost any plasmid DNA could be employed as substrate, it is now customary to use the small (4.3-kb) plasmid pBR322, thereby maximizing the electrophoretic separation of relaxed and supercoiled DNA. Topo IV activity is assayed by following its ATP-dependent decatenation of kinetoplast DNA (kDNA) again using agarose gel electrophoresis. These are the assays of choice in investigating the effects of catalytic inhibitors such as coumarins. By contrast,

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a DNA cleavage assay is used for quinolones and other topoisomerase II poisons that act by stabilizing the topoisomerase cleavage complex on DNA resulting in double-stranded DNA breaks. Given the recent recognition that either gyrase or topo IV may be the potential target (14,15), it is now usual to test inhibitor effects on both enzymes. The following sections describe the gyrase supercoiling assay, the topoisomerase IV decatenation assay, and a DNA cleavage assay applicable to both gyrase and topo IV. 2. Materials 2.1. DNA Supercoiling Assay 1. 3X Gyrase assay solution: 105 mM Tris-HCl, pH 7.5, 18 mM MgCl2 , 5.4 mM spermidine, 72 mM KCl, 15 mM DTT, 1.08 mg/mL BSA, 19.5% glycerol (w/v), and (optional) 90 μg/mL E. coli tRNA (Calbiochem). 2. Gyrase dilution buffer: 50 mM Tris-HCl, pH 7.5, 0.2 M KCl, 5 mM DTT, 1 mM EDTA, 3 mg/mL BSA, 50% glycerol. (3X assay solution and dilution buffer can be made up and stored in aliquots at –20 °C). 3. 50-mM ATP solution: 27.5 mg ATP (disodium salt) dissolved in 1 mL 100 mM NaOH (see Note 1). 4. Relaxed pBR322 DNA (see Note 2). 5. 5X dye mix: 5% SDS, 25 % glycerol, 0.25 mg/ml bromophenol blue. 6. Agarose gel: 1% agarose in TBE (90 mM Tris base, 90 mM boric acid, 2.5 mM EDTA). 7. 10 mg/mL ethidium bromide (EtBr) (see Note 3). 8. Gyrase preparation or the individual purified GyrA and GyrB subunits. 9. Prospective gyrase inhibitor and known control inhibitor. 10. Gel documentation system, e.g., Alpha Innotech digital camera and associated software. Such instrumentation allows quantification of DNA bands and consequently has largely superseded image capture on Polaroid film using a Land camera and uv transilluminator.

2.2. Decatenation Assay for Topo IV 1. 4X Decatenation buffer: 160 mM Tris-HCl, pH 7.5, 24 mM MgCl2 , 40 mM DTT, 40 mM NaCl, 800 mM potassium glutamate, and 0.2 mg/mL BSA. 2. 1X Decatenation buffer. 3. 50 mM ATP. 4. Kinetoplast DNA: supplied by Topogen, Inc. 5. 5X Dye mix, as in previous section. 6. Topo IV or individual ParC and ParE subunits. 7. Agarose gel: 1% in TBE (90 mM Tris base, 90 mM boric acid, 2.5 mM EDTA). 8. 10 mg/mL EtBr (see Note 3). 9. Gel documentation system, as in previous section.

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2.3. Drug-Promoted DNA Cleavage by Gyrase or Topo IV 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

3X DNA gyrase buffer (or 4X topo IV buffer). Supercoiled pBR322 supplied by New England Biolabs. Linear pBR322 DNA. Sodium dodecyl sulfate. Proteinase K. 5X Dye mix. Gyrase (GyrA and GyrB subunits) or topo IV (ParC and ParE subunits). Agarose gel: 1% in TBE. 10 mg/mL EtBr (see Note 3). Gel documentation system, as in previous section.

3. Methods 3.1. DNA Supercoiling Assay This section describes how to measure and titrate the ATP-dependent DNA supercoiling activity of gyrase prior to testing the effect of inhibitors on the enzyme reaction. The assay uses relaxed pBR322 DNA as substrate and in principle may employ gyrase activity present in bacterial extracts, native enzyme purified from bacteria, or individual GyrA and GyrB proteins obtained from bacterial extracts by affinity chromatography on novobiocin-Sepharose (17). Crude extracts from bacteria should be avoided as they contain large amounts of nucleases that lead to nicking of plasmid DNA and thereby abrogate the assay. It is better to purify the enzyme by ammonium sulfate and Polymin P fractionation, plus column chromatography, including tRNA when assaying to suppress any nuclease activity. Highly purified DNA gyrases from E. coli and Micrococcus luteus are available from John Innes Enterprises and from Gibco BRL. Increasingly, recombinant His-tagged GyrA and GyrB proteins are being prepared for a variety of bacterial species using E. coli strains bearing the appropriate gyrA or gyrB genes cloned in inducible vectors such as pET (3–5). Although construction of expression plasmids is time-consuming, it has the advantage that the tagged GyrA and GyrB proteins can be recovered in high purity (>95% homogeneity) from E. coli extracts in a single step by nickel chelate chromatography (4,5). Neither subunit alone is active, but gyrase activity can be reconstituted by mixing GyrA and GyrB. Each subunit is assayed in the presence of an excess of the complementing protein so that the specific activity of each subunit can be measured individually. This approach, based on Ref. 4, is described using S. pneumoniae GyrA and GyrB proteins.

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1. Gyrase assays are performed in 35 mM Tris-HCl, pH 7.5, 24 mM KCl, 6 mM MgCl2 , 1.8 mM spermidine, 0.36 mg/mL bovine serum albumin (BSA), 1.4 mM ATP, 0.4 μg relaxed pBR322, and various dilutions of the gyrase activity to be measured (final volume 35 μL). When using individually purified GyrA and GyrB proteins, the subunit to be assayed is diluted into gyrase dilution buffer (see Note 4) (on ice) and added to gyrase assay buffer on ice containing an excess (>10U) of the complementing subunit. 2. Make up a cocktail on ice containing 11.7 μL 3X gyrase assay mix, 2 μL of relaxed DNA (i.e., 0.4 μg), ATP, complementing subunit (where needed), and sterile water to 33 μL. These quantities should be increased according to the number of assays to be run. The final mix is distributed to 1.5-mL Eppendorf tubes on ice, and the complementing gyrase subunit (2 μL) is added. To mix the reactants, gently tap the tube or withdraw and eject the solution several times into the tip of a Gilson pippetor. Transfer the tubes to a circulating water bath at 25 °C and incubate for 1 h. 3. Stop the reaction by adding 8 μL of 5X dye mix and separate the DNA products by electrophoresis on a 1% agarose gel run overnight at 2 V/cm (see Note 5). The gel should be stained in TBE containing 0.5 μg/mL EtBr for 30 min to 1 h, destained for 1 h in TBE, and photographed under UV irradiation using a gel documentation system, e.g., the Alpha Innotec digital camera and associated software. 4. One unit of gyrase activity is defined as the amount of enzyme that supercoils 50% of the relaxed pBR322 substrate under these reaction conditions. 5. Figure 1 shows an assay for gyrase activity wherein an appropriately diluted S. pneumoniae GyrA subunit is assayed in the presence of excess GyrB, and vice versa. The amount of GyrA and GyrB that converts 50% of the input DNA to the supercoiled form is in each case 15 ng, corresponding to specific activities of 6.7 × 104 U/mg for both subunits (see Note 6). Similar conditions can be used to assay gyrase subunits from other bacterial species. 6. Figure 2 illustrates the use of the assay to investigate the effects of ciprofloxacin (see Note 7) on supercoiling activity. No more than 2 U of gyrase activity are used per assay (sufficient to just supercoil all the input DNA) so that the reaction is optimally sensitive to any inhibitory effects. For ciprofloxacin, the drug concentration that inhibits DNA supercoiling by 50% (the IC50 ) is 40 μM (Fig. 2).

3.2. kDNA Decatenation Assay for Topoisomerase IV Topo IV is conveniently assayed by its ability to decatenate kDNA, a network of topologically linked DNA circles made up of several thousand 2-kb minicircles and a few larger maxicircles (4). Reaction products are analyzed by agarose gel electrophoresis, exploiting the fact that released minicircles migrate into the gel, whereas the kDNA network is too large to move into the agarose and is retained in the wells. kDNA from Crithidia fasciculata is available commercially from Topogen. Topo IV from E. coli can be obtained from

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N R S

Fig. 1. Reconstitution and titration of DNA supercoiling activity using S. pneumoniae GyrA and GyrB subunits. Relaxed pBR322 substrate was mixed with either 300 ng GyrB (lanes 2 to 7) or 100 ng GyrA (lanes 8 to 13). To titrate GyrA activity, the reaction mix containing 1 mM ATP was incubated with 60, 30, 15, 7.5, 3.75, and 1.87 ng GyrA protein (lanes 2–7), respectively. Similarly, to assay GyrB, 120, 60, 30, 15, 7.5, and 3.75 ng GyrB was added to lanes 8 to 13, respectively. After incubation, products were resolved by electrophoresis on a 1% agarose gel. Lane 1, relaxed pBR322 control, which contains a small amount of nicked DNA. N, R, and S denote the nicked, relaxed, and supercoiled pBR322, respectively. DNA bands migrating between N and S are partially supercoiled DNA products.

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Fig. 2. Inhibition of DNA supercoiling by ciprofloxacin. Relaxed pBR322 DNA was incubated with S. pneumoniae gyrase (2U) in the absence (lane 3) or presence of ciprofloxacin at 10, 20, 40, 80, and 160 μM (lanes 4 to 8). Increasing drug levels lead to dose-dependent inhibition of DNA supercoiling. Lane 1, supercoiled pBR322 control; lane 2, relaxed pBR322 control.

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John Innes Enterprises. Recombinant topo IV from a number of Gram-positive pathogens has been reconstituted from subunits purified from overexpressing parC and parE strains. Prior to inhibitor studies, it is necessary to titrate topo IV activity to ascertain the correct amount of enzyme to use. Where individual ParC and ParE subunits are available, each protein is assayed in the presence of an excess of the other, allowing its specific activity to be assessed. Topo IV subunits are combined accordingly for inhibitor studies. 1. Topo IV assays are carried out in a decatenation buffer [40 mM Tris-HCl, pH 7.5, 6 mM MgCl2 , 10 mM NaCl, 10 mM DTT, 50 μg/mL BSA, and 200 mM potassium glutamate, to which are added 1 mM ATP, 450 ng of kDNA, and various amounts of the topo IV, ParC, or ParE subunit (diluted into 1X decatenation buffer) (final volume 20 μL)]. 2. A cocktail is made up on ice using 4X topo IV assay mix, ATP, kDNA (2 μL, i.e., 450 ng), ParE or ParC subunit (where appropriate), and sterile water to 18 μL. These quantities are multiplied according to the number of assays to be run. The solution is distributed in 18-μL aliquots to 1.5-mL Eppendorf tubes on ice and complementing the topo IV subunit added (2 μL). Reaction mixtures are incubated in a circulating water bath at 37 °C for 1 h. 3. The reaction is terminated by addition of 5X dye mix (4 μL). DNA products are separated by electrophoresis on a 1% agarose gel run at 2 V/cm overnight. After staining with EtBr, the gel is photographed under UV light on a gel documentation system (for details, see Sections 2.1 and 3.1). 4. One unit of topo IV activity can be defined as the amount of enzyme that decatenates 50% of the input kDNA under these reaction conditions. 5. Figure 3 shows a representative assay for decatenation activity in which various amounts of an S. pneumoniae ParC and ParE are assayed in the presence of excess complementing subunit (see Note 8). Under these conditions, 0.45 ng of ParC and 6.25 ng of ParE were sufficient to give 50% decatenation, corresponding to specific decatenation activities of 2 × 106 U/mg and 1.5 × 105 U/mg, respectively. 6. Figure 4 demonstrates the inhibition of topo IV decatenation activity by ciprofloxacin. The amount of topo IV is sufficient to just decatenate 0.45 μg of kDNA. For ciprofloxacin, the IC50 (the amount of drug that gives 50% inhibition of decatenation) is 10 μM (Fig. 4, lane 6).

3.3. DNA Cleavage Assays for Gyrase and Topo IV Some topoisomerase inhibitors such as quinolones exert their bactericidal effects by stabilizing the cleavage complex of gyrase or topo IV on DNA (1,4,12). Cellular processes in vivo or protein denaturation in vitro can convert the complex into a frank double-stranded DNA break that is thought to be the lethal lesion. The assay for cleavage complex stabilization involves measuring

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Fig. 3. Titration of DNA decatenation activity of S. pneumoniae topo IV. Kinetoplast DNA was incubated with ATP and either excess ParE (100 ng) and ParC at 30, 15, 7.5, 3.75, 1.8, 0.9, 0.45 ng, respectively (lanes 2–8), or excess ParC (30 ng) and ParE at 100, 50, 25, 12.5, 6.25, 3.1, and 1.5 ng, respectively (lanes 9–15). Reactions were stopped and products were resolved by electrophoresis in 1% agarose gel. Lane 1, kDNA control. Bands with electrophoretic mobility between that of kDNA and released minicircles (monomer) are partially decatenated DNA intermediates.

the ability of the inhibitor to promote linearization of pBR322 DNA mediated by gyrase or by topo IV (4). Supercoiled pBR322 DNA is incubated with enzyme in the presence or absence of inhibitor. The detergent sodium dodecyl sulfate (SDS) is added to denature the cleavage complex and thereby release broken

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Fig. 4. Inhibition of S. pneumoniae topo IV decatenation activity by ciprofloxacin. Kinetoplast DNA was incubated with S. pneumoniae topo IV (2U) in the presence of ciprofloxacin at 0, 1, 2.5, 5, 10, 20, and 40 μM, respectively (lanes 2–8). Lane 1, kDNA control.

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DNA, and following proteinase K digestion (to remove GyrA or ParC subunit linked to DNA), the DNA products are examined by agarose gel electrophoresis. Trapping of the cleavage complex is revealed by the generation of linear pBR322 DNA. It should be noted that detection of DNA cleavage requires the use of stoichiometric amounts of DNA substrate and gyrase or topo IV, i.e., typically 50–100 times more enzyme than employed in assays of catalytic function. The following protocol has been developed for S. pneumoniae gyrase and topo IV but is widely applicable to the enzymes from other bacteria. 1. DNA cleavage assays for gyrase are performed in the absence and presence of inhibitor in 35 mM Tris-HCl, pH 7.5, 24 mM KCl, 6 mM MgCl2 , 1.8 mM spermidine, 0.36 mg/mL bovine serum albumin (BSA), supercoiled pBR322, and DNA gyrase (final volume 20 μL). Except for the higher enzyme levels, these conditions are similar to those used in the gyrase supercoiling assay (see Sections 2.1 and 3.1), except ATP is omitted (see Note 9). Cleavage of supercoiled pBR322 by topo IV is also observed under these gyrase conditions (allowing direct comparison) but is more efficient using a buffer containing 40 mM Tris-HCl, pH 7.5, 6 mM MgCl2 , 10 mM NaCl, 10 mM DTT, 50 μg/mL BSA, and 200 mM potassium glutamate, i.e., similar to the decatenation assay conditions. 2. Make a reaction mix on ice containing 6.7 μL 3X gyrase cleavage buffer or 5 μL of 4X topo IV cleavage buffer, supercoiled pBR322 DNA (0.4 μg), and sterile water to 16 μL. A multiple of these quantities can be made up depending on the number of assays to be run. The mix is distributed into 1.5-mL Eppendorf tubes on ice containing various concentrations of the inhibitor (in 2 μL) to be assayed. Reaction is initiated by adding 0.45 μg GyrA (or ParC) and 1.7 μg GyrB (or ParE) (in 2 μL) that had been mixed and pre-incubated on ice for 10 min. After mixing, the tubes are incubated in a circulating water bath at 25 °C for 1 h. 3. Induce DNA cleavage by adding 3 μL of 2% (w/v) SDS to each tube taken directly from the water bath and vortex. Add 3 μL of 1 mg/mL proteinase K (see Note 10), vortex, and incubate for 30 min at 37 °C. 4. Reactions are stopped by adding 7 μL of 5X dye mix, and the DNA products are separated by electrophoresis in 1% agarose. The gel is stained with EtBr and photographed as described in earlier sections. 5. Figure 5 shows a representative DNA cleavage experiment for gyrase and topo IV. DNA cleavage can be quantified in terms of the CC25 , the drug concentration that converts 25% of the input DNA to the linear form (see Notes 11 and 12). The ciprofloxacin CC25 values for gyrase and topo IV are 80 μM and 5–10 μM, respectively (see Note 13). Quinolone CC25 values (and indeed the IC50 values) are elevated at least 8- to 16-fold for S. pneumoniae gyrase and topo IV complexes reconstituted with GyrA and ParC subunits bearing quinolone resistance mutations of Ser79Phe and Ser81Phe, respectively (18).

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Fig. 5. DNA cleavage induced by S. pneumoniae gyrase (A) and topo IV (B) in the presence of ciprofloxacin. Supercoiled pBR322 was incubated (A) with gyrase (0.45 μg GyrA and 1.7 μg GyrB) and ciprofloxacin at 0, 20, 40, 80, 160, and 320 μM (lanes 3–8, respectively) or (B) topo IV (0.45 μg ParC and 1.7 μg ParE) and ciprofloxacin at 0, 2.5, 5, 10, 20, and 40 μM (lanes 3–8, respectively). DNA cleavage was induced with SDS, and after proteinase K digestion, DNA products were displayed by 1% agarose gel electrophoresis. Lane 1, supercoiled pBR322 control; lane 2, linear pBR322 control. N, L, and S denote nicked, linear, and supercoiled pBR322, respectively.

4. Notes 1. ATP solutions should be stored as aliquots at –20 °C and discarded after use. 2. Relaxed pBR322 may be purchased from John Innes Enterprises. Alternatively, it can be readily prepared by relaxation of supercoiled pBR322 (New England Biolabs) using either human topoisomerase I (from Topogen) or the calf thymus topo I from New England Biolabs according to the protocol detailed in Ref 19. 3. A note of caution: EtBr is a mutagen. Gloves should be worn when handling EtBr solutions, and both should be disposed of using proper procedures. 4. Do not add more than 2 μL of gyrase diluent to the assay: It contains a high concentration of salt and can inhibit the reaction. 5. For optimal separation of relaxed and supercoiled DNA, gels should be run slowly overnight. EtBr should not be included in the agarose gel as it intercalates into DNA, changing the mobility of relaxed DNA to that of supercoiled DNA. 6. Relaxed DNA prepared by treatment of supercoiled DNA with a eukaryotic topoisomerase I is made up of a Gaussian distribution of topoisomers bearing differing linking numbers, which are resolved in the gel as a discrete ladder of bands (see Fig. 1, lane 1). Preparations of supercoiled DNA normally contain a small amount of nicked DNA. Any nuclease contamination in the gyrase preparation converts the relaxed DNA ladder to nicked DNA that migrates as a single band and cannot be supercoiled by gyrase. 7. Ciprofloxacin is a quinolone inhibitor of gyrase/topo IV. It exerts its inhibitory action by interfering with DNA resealing in the cleavage complex. 8. Topo IV has both decatenation and DNA relaxation activities. Therefore, the released minicircles migrate as relaxed DNA species.

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9. Inclusion of ATP has no significant effect on the level of quinolone-induced DNA cleavage mediated by the pneumococcal gyrase and topo IV. However, ATP does enhance the efficiency of DNA cleavage by other type II topoisomerases, e.g., E. coli gyrase. 10. The solution of proteinase K in sterile water can be stored frozen as aliquots at –20 °C for several months. 11. Linear pBR322 DNA used as the control in Fig. 5 is made by digestion of supercoiled pBR322 with the restriction enzyme EcoRI. 12. In the absence of quinolone, the DNA breakage-resealing equilibrium for gyrase and topo IV is over to the side of sealed DNA, presumably as a safeguard against inadvertent DNA breakage in vivo. Consequently, little or no linear DNA is detectable for gyrase in the absence of quinolone (Fig. 5A, lane 3). However, for topo IV, more cleavage complex appears to be present at equilibrium, and we consistently observe that topo IV produces some linear DNA, albeit at a low level, in the absence of drug (Fig. 5B, lane 3). 13. DNA cleavage is a stoichiometric reaction, i.e., the more enzyme that is added, the greater the amount of cleavage at any given drug concentration. The levels of gyrase/topo IV used in the assay have been chosen so that enzyme is in excess of DNA substrate, producing a proper dose response as drug concentrations are increased. Full-length linear pBR322 product arises from DNA cleavage by a single enzyme cleavage complex per DNA molecule. At higher drug levels, multiple enzyme complexes are trapped on each DNA molecule, generating substantial DNA fragmentation, seen as a smear of DNA products below the linear band (Fig. 5B, lanes 7 and 8).

Acknowledgments X.-S. Pan and this work were supported by Project Grant BBD01882X1 from the Biotechnology and Biological Sciences Council of the United Kingdom. References 1 Drlica, K., and Zhao, X. (1997) DNA gyrase, topoisomerase IV, and the 41. quinolones. Microbiol. Mol. Biol. Rev. 61, 377–392. 2 Gellert, M., Mizuuchi, K., O’Dea, M. H., and Nash, H. A. (1976) DNA gyrase:An 2. enzyme that introduces superhelical turns into DNA. Proc. Natl. Acad. Sci.USA 73, 3872–3876. 3 Mizuuchi, K., Mizuuchi, M., O’Dea, M. H., and Gellert, M. (1984) Cloning and 3. simplified purification of Escherichia coli DNA gyrase A and B proteins. J. Biol. Chem. 259, 9199–9201. 4 Pan, X.-S., and Fisher, L. M. (1999) Streptococcus pneumoniae DNA gyrase 4. and topoisomerase IV: Overexpression, purification, and differential inhibition by fluoroquinolones. Antimicrob. Agents Chemother. 43, 1129–1136.

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5 Aubry, A., Veziris, N., Cambau, E., Truffot-Pernot, C., Jarlier, V., and Fisher, L. M. 5. (2006) Novel gyrase mutations in quinolone-resistant and -hypersusceptible clinical isolates of Mycobacterium tuberculosis: Functional analysis of mutant enzymes. Antimicrob. Agents Chemother. 50, 104–112. 6 Mizuuchi, M., Fisher, L. M., O’Dea, M. H., and Gellert, M. (1980) DNA gyrase 6. action involves the introduction of transient double strand breaks into DNA. Proc. Natl. Acad. Sci. USA. 77, 1847–1851. 7 Corbett, K. D., and Berger, J. M. (2004) Structure, molecular mechanisms, and 7. evolutionary relationships in DNA topoisomerases. Ann. Rev. Biophys. Biomol. Struct. 33, 95–118. 8 Corbett, K. D., Schoeffler A. J., Thomsen, N. D., and Berger, J. M. (2005) The 8. structural basis for substrate specificity in DNA topoisomerase IV. J. Mol. Biol. 351, 545–561. 9 Kato, J., Nishimura, Y., Imamura, R., Niki, H., Higara, S., and Suzuki, S. (1990) 9. New topoisomerase essential for chromosome segregation in E. coli. Cell 63, 393–404. 10 Maxwell, A., and Lawson, D. M. (2003) The ATP binding site of type II topoiso10. merases as a target of antibacterial drugs. Curr. Top. Med. Chem. 3, 283– 303. 11 Oram, M., Dosanjh, B., Gormley, N. A., Smith, C. V., Fisher, L. M., Maxwell, 11. A., and Duncan, K. (1996) The mode of action of GR122222X, a novel inhibitor of DNA gyrase. Antimicrob. Agents Chemother. 40, 473–476. 12 Drlica, K., and Malik, M. (2003) Fluoroquinolones: Action and resistance. Curr. 12. Top. Med. Chem. 3, 249–282. 13 Grossman, T. H., Bartels, D. J., Mullin, S., Gross, C. H., Parsons, J. D., Liao, Y., 13. Grillot, A.L., Stamos, D., Olson, E. R., Charifson, P. S., and Mani, N. (2007) Dual targeting of GyrB and ParE by a novel aminobenzimidazole class of antibacterial compounds. Antimicrob. Agents Chemother. 51, 657–666. 14 Pan, X.-S., and Fisher, L. M. (1997) Targeting of DNA gyrase in Streptococcus 14. pneumoniae by sparfloxacin: Selective targeting of gyrase or topoisomerase IV by quinolones. Antimicrob. Agents Chemother. 41, 471–474. 15 Pan, X.-S., and Fisher, L. M. (1998) DNA gyrase and topoisomerase IV are dual 15. targets of clinafloxacin action in Streptococcus pneumoniae. Antimicrob. Agents Chemother. 42, 2810–2816. 16 Zhanel, G. G., Fontaine, S., Adam, H., Schurek, K., Mayer, M., Noreddin, A. M., 16. Gin, A. S., Rubinstein, E., and Hoban, D. J. (2006) A review of new fluoroquinolones: Focus on their use in respiratory tract infections. Treatment Respir. Med. 5, 437–465. 17 Staudenbauer, W. L., and Orr, E. (1982) DNA gyrase: Affinity chromatog17. raphy on novobiocin-Sepharose and catalytic properties. Nucleic Acids Res. 9, 3589–3603. 18 Pan, X.-S., Yague, G., and Fisher, L. M. (2001) Quinolone resistance mutations 18. in Streptococcus pneumoniae GyrA and ParC proteins: Mechanistic insights into

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quinolone action from enzymatic analysis, intracellular levels, and phenotypes of wild-type and mutant proteins. Antimicrob. Agents Chemother. 45, 3140–3147. 19 Sayer, P. J., Goble, M. L., Oram, M., and Fisher, L. M. (2001) Plasmid supercoiling 19. by DNA gyrase. In Methods in Molecular Biology, Vol. 95: DNA Topoisomerase Protocols, Part II: Enzymology and Drugs (N. Osheroff and M. A. Bjornsti, eds.), Humana Press Inc, Totowa, NJ, pp. 25–33.

3 A Method to Assay Inhibitors of DNA Polymerase IIIC Activity Michelle M. Butler and George E. Wright

Summary The need for new drugs to treat infections caused by antibiotic-resistant bacterial strains has prompted many studies to identify novel targets in pathogenic bacteria. Among the three DNA polymerases expressed by bacteria, one of these, designated pol III, is responsible for DNA replication and growth of bacteria and, therefore, warrants consideration as a drug target. However, the pol III enzymes of Gram-positive and Gram-negative species are quite different, and the Gram-positive enzyme pol IIIC has been more extensively studied as a drug target than the Gram-negative enzyme pol IIIE. DNA polymerases are unique enzymes with respect to the five substrates (four dNTPs, one of which is radiolabeled, and primer:template DNA) that they typically utilize. Variations of the assay, e.g., by leaving out one dNTP but allowing measurable incorporation of the remaining substrates, or use of homopolymer primer:templates, may be used to simplify the assay or to obtain mechanistic information about inhibitors. Use of gel analysis of primer extension assays can also be applied to study alternate substrates of DNA polymerases. Methods to isolate pol IIIC from Gram-positive bacterial cells and to clone and express the polC gene are described in this chapter. In addition, the assay conditions commonly used to identify and study the mechanism of inhibitors of pol IIIC are emphasized.

Key Words: DNA polymerase; pol IIIC; pol IIIE; isolation; cloning; DNA polymerase assay.

1. Introduction Bacteria contain three distinct DNA polymerases, designated pol I, pol II, and pol III (1). Pol III has been found to be responsible for DNA replication and growth of bacteria and is, therefore, the most relevant enzyme as a drug From: Methods in Molecular Medicine, Vol. 142: New Antibiotic Targets Edited by: W. Scott Champney © Humana Press Inc., Totowa, NJ

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target. However, the pol III enzymes of Gram-positive and Gram-negative species differ considerably. The Gram-positive enzyme, now designated pol IIIC, is a high-molecular-weight (MW) protein with polymerase and 3’,5’- and 5’,3’-exonuclease activities and has been exploited by several groups as a drug target (2–5). The Gram-negative enzyme, designated pol IIIE, is smaller and contains only polymerase activity. Recently, the pol II species from Grampositive bacteria has been renamed pol IIIE (because of its similarity to the corresponding enzyme from Gram-negatives), and it too has been shown to have an essential role in DNA replication (6). Competitive inhibitors of Gram-positive pol IIIC have been studied for some time and were, in fact, instrumental in defining the role of pol IIIC in bacterial DNA replication. In addition, study of the mechanism, potency, and selectivity of pol IIIC inhibitors has resulted in compounds currently under investigation as antibacterial agents (2,5). DNA polymerases are unique enzymes with respect to the five substrates (four dNTPs and primer:template DNA) that they typically utilize. The finding that one dNTP can be removed from the assay (“truncated assay”), but measurable incorporation of the remaining substrates is permitted, forms the basis of a powerful, direct measure of the Ki of an inhibitor competitive with the removed dNTP (7). Alternately, homopolymer primer:templates utilizing one dNTP can be used to great effect in defining both the potency and competitive nature of inhibition by novel compounds. The assay conditions commonly used to identify and study the mechanism of inhibitors of pol IIIC are emphasized in this article, and both isolation of pol IIIC from bacterial cells and its cloning and expression are described. 2. Materials 2.1. Preparation of B. subtilis Pol IIIC 2.1.1. Isolation of Native B. subtilis Pol IIIC 1. 2. 3. 4. 5. 6. 7. 8. 9.

B. subtilis NB841, obtained from Dr. Neal Brown. 20 mM Tris-acetate, pH 8.2, 10 mM magnesium acetate, 0.5 mM EDTA. 15 mM potassium phosphate, pH 7.4, 300 mM ammonium sulfate. 10 mM potassium phosphate, pH 7.4, 200 mM ammonium sulfate. 10 mM potassium phosphate, pH 6.5. Hydroxylapatite (Clarkson Chemical Co.). DEAE cellulose DE 52 (Whatman). Sephadex G25, G200 (Pharmacia). Agarose (Bio-Rad).

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10. Denatured DNA cellulose, prepared as described by Alberts et al. (8). 11. Triton X-100 (Sigma).

2.1.2. Isolation of Recombinant B. subtilis Pol IIIC 1. Plasmid pSGA04, obtained from Dr. John Lowenstein. 2. Restriction enzymes, HpaI, XhoI, BamHI, and EcoRI as well as E. coli. Klenow pol I can be purchased from New England Biolabs and is stored at –20 °C. 3. LB; Luria broth, Lennox (Difco). 4. PBS: 150 mM NaCl, 50 mM potassium phosphate, pH 7.5. 5. Column buffer: 50 mM potassium phosphate, pH 7.5, 20% glycerol, 1 mM PMSF, 2 mM -mercaptoethanol. 6. IMAC agarose affinity gel (Sigma-Aldrich) is stored at 4 °C. 7. MacroPrep High Q (Bio-Rad) is stored at 4 °C. 8. MacroPrep column buffer: 50 mM potassium phosphate, pH 7.5, 10% glycerol, 5 mM PMSF, 2 mM -mercaptoethanol, stored at 4 °C.

2.2. DNA Substrates 2.2.1. Activated Calf Thymus DNA 1. Calf thymus DNA (Worthington Biochemical Corporation) is stored at 4 °C. 2. 50 mM Tris-HCl, pH 7.5, sterile-filtered into a sterile flask. 3. Deoxyribonuclease I (DNAse, Sigma), dissolved in dH2 O at 5 mg/mL and stored at –20 °C. 4. Deoxyribonucleotides: dATP, dCTP, dGTP, dTTP (100 mM stock solutions, Fermentas). 5. [Methyl-3 H]-thymidine 5’-triphosphate, ammonium salt (Amersham/GE Healthcare). 6. DNA-free assay mix: 50 mM Tris-HCl, pH 7.5 (30 mM final), 16.67 mM magnesium acetate (10 mM final), 6.67 mM dithiothreitol (4 mM final), 26.67% (v/v) glycerol (16% final), 41.6 μM dATP, dCTP, dGTP (25 μM final), 16.67 μM [3 H] TTP, 2 μCi/nmole (10μM, 1.2 μCi/nmole final). This mixture can be stored at –20 °C for several years.

2.2.2. Synthetic Primer:Templates 1. 2. 3. 4. 5.

Oligodeoxythymidylate, (dT)12−18 (Midland Certified Reagent Company). Polydeoxyadenylic acid, poly(dA) (Midland Certified Reagent Company). Oligodeoxyguanylate (dG)12−18 (Midland Certified Reagent Company). Polydeoxycytidylic acid, poly(dC) (Midland Certified Reagent Company). Powders and solutions of polymers and oligonucleotides should be stored at –20 °C.

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2.3. Pol IIIC Assays 2.3.1. Standard Pol IIIC Assay 1. 96-well assay plate (U-bottom polypropylene, AbGene). 2. Chemicals such as Tris-HCl, magnesium acetate, glycerol, dithiothreitol (DTT), trichloroacetic acid (TCA), hydrochloric acid (HCl), sodium pyrophosphate, and ethanol can be purchased from Sigma-Aldrich or VWR Scientific. 3. Deoxyribonucleotides: dATP, dCTP, dGTP, dTTP (100 mM stock solutions, Fermentas). 4. [Methyl-3 H]-thymidine 5’-triphosphate, ammonium salt (Amersham/GE Healthcare). 5. Assay glass fiber filter plates (96-well, Multiscreen-HTS FC Barex, Millipore). 6. Liquid scintillation cocktail (OptiPhase Supermix, Perkin Elmer).

2.3.2. Pol IIIC Assay Using Synthetic Template:Primers 1. [Methyl-3 H]-thymidine 5’-triphosphate, ammonium salt, and Deoxy-[8-3 H]guanosine 5’-triphosphate, ammonium salt (Amersham/GE Healthcare).

2.3.3. Primer Extension Assay 1. The primer is a 17mer, 5’-GTAAAACGACGGCCAGT-3’ (M13/pUC-20 sequencing primer, New England Biolabs). 2. The template is a custom-synthesized 29mer 3‘CATTTTGCTGCCGGTCACATGCCGATCCC-5’ (Operon). 3. T4 polynucleotide kinase (New England Biolabs). 4. [32 P]-ATP, triethylammonium salt (Amersham/GE Healthcare). 5. Calf thymus terminal deoxyribonucleotidyl transferase (New England Biolabs). 6. Spermidine, Sephadex G-25 (Sigma-Aldrich). 7. Deoxyribonucleotide dGTP (100 mM stock solution, Fermentas). 8. Extension buffer: 20 mM potassium phosphate, pH 7.5, 8 mM MgCl2 , 4 mM DTT, 0.1 mM EDTA. 9. Stop buffer: 95% formamide, 20 mM EDTA, 0.05% bromphenol blue, and 0.05% xylene cyanol FF.

3. Methods 3.1. Preparation of B. subtilis Pol IIIC 3.1.1. Isolation of Native B. subtilis Pol IIIC Purification of native pol III from B. subtilis has been described, starting with a strain that is deficient in pol I (9,10). 1. All buffers contain 20% (v/v) glycerol and 5 mM -mercaptoethanol, and purification procedures are done at 0–4 °C. Protein concentration is estimated by the

A Method to Assay Inhibitors of DNA Polymerase IIIC Activity

2.

3.

4.

5.

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method of Lowry et al. (11). Five g of packed B. subtilis NB841 are suspended in 10 mL of a buffer consisting of 20 mM Tris-acetate, pH 8.2, 10 mM magnesium acetate, and 0.5 mM EDTA, and ruptured in a French pressure cell. The lysate is freed of debris by centrifugation at 17,000X g for 30 min, and the supernatant is further centrifuged for 3 h in a Spinco 50.1 rotor at 42,000 rpm (200,000X g). The high-speed supernatant (10 mL; 20 mg of protein per mL) is diluted with 20 mL of 16 mM potassium phosphate buffer, pH 7, containing 300 mM ammonium sulfate, and passed at a rate of 0.5 mL/min through a 13 × 1.65-cm column of DEAE-cellulose equilibrated with 10 mM potassium phosphate buffer (pH 7.4) containing 200 mM ammonium sulfate. The first 15 mL of eluant is discarded; the subsequent 45 mL, containing 85% of input protein, is mixed with 22 g of ground ammonium sulfate and stirred for 90 min. The precipitated protein is harvested by centrifugation, dissolved in 6 mL of 10 mM potassium phosphate, pH 6.5, and freed of ammonium sulfate by passage through Sephadex G-25 in the same buffer (Fraction IV). All of the solutions contain 20% glycerol, 10 mM magnesium acetate, 25 mM -mercaptoethanol, 0.5% Triton X-100, and potassium phosphate, pH 6.5 (referred to as phosphate) at a specified concentration. Fraction IV is diluted to 2 mg per mL in 10 mM phosphate and applied to a 100 mL column (2 × 32 cm) of denatured DNA-cellulose equilibrated with the same buffer. The column is eluted with a linear gradient (0.1 to 0.4 M; volume 1000 mL; 3.3 mL per min) of phosphate. The polymerase activity, which emerges as a single peak between 0.22 and 0.29 M phosphate, is pooled in a volume of 190 mL, diluted 4-fold with 10 mM phosphate, and applied to a 5 mL column (1.1 × 5 cm) of DEAE-cellulose washed with 10 mM phosphate. The enzyme, which is retained on the column, is eluted with 0.5 M phosphate in a volume of 6.5 mL (Fraction V). Fraction V is diluted 7-fold with 10 mM phosphate and applied to a 25 mL column (1.5 × 15 cm) of hydroxylapatite equilibrated with 10 mM phosphate, and eluted with a linear gradient (0.05 to 0.3 M; volume, 600 mL; 2 mL per min) of phosphate. Enzyme activity, which emerges as a single peak between 0.15 and 0.19 M phosphate, is pooled in a volume of 96 mL and diluted 3-fold with 10 mM phosphate. The pool is applied to a l mL column (0.56 × 4 cm) of DEAE-cellulose washed with 10 mM phosphate, and the enzyme is eluted in a volume of 0.8 mL of 0.5 M phosphate (Fraction VI). Fraction VI is applied to a column (1.1 × 54 cm) of Sephadex G-200 equilibrated with 0.5 M phosphate and eluted at a rate of 0.9 mL per hour. The enzyme emerges as a symmetrical peak approximately 5 mL after the void volume (15 mL, blue dextran 2000); 80% of the activity is pooled in a volume of 5 mL, diluted 7-fold with 10 mM phosphate, and concentrated into 0.25 mL of 0.5 M phosphate with a 0.25 mL column (0.56 × 1 cm) of DEAE-cellulose (Fraction VII).

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3.1.2. Cloning, Overexpression, and Isolation of Recombinant B. subtilis Pol IIIC 1. Engineering of B. subtilis polC into expression plasmid pSGA04 (12) requires the following five steps that have been described elsewhere (13): (i) introduction, with PCR, of a new HpaI restriction site at nucleotide 16 of a form of Bs polC that has been engineered previously to contain an XhoI site at position 1246 (14); (ii) excision of the resulting 1228-bp HpaI-XhoI fragment; (iii) recloning of the corresponding DNA into a fully wild-type Bs polC in the vector pKC30(15); (iv) excision of the polC gene lacking the first 15 bases from pKC30 as an HpaI-BamHI fragment; and (v) inserting the fragment into pSGA04, which has been digested with EcoRI, filled in by treatment with Klenow pol I, and subsequently digested with BamH1to create expression vector pSGA04-polC (see Note 1). 2. Plasmid pSGAO4-polC is introduced into E. coli SG101 by transformation, and transformant colonies are grown at 30 °C to an A600 of approximately 1 in LB expression medium containing 15 μg/mL kanamycin as described (13). The culture is then chilled on ice to a temperature of 15 °C, IPTG is added to a concentration of 1 mM to induce expression of pSGAO4-polC, and incubation at 15 °C is continued, with shaking, for 18 h. 3. The cells are harvested by centrifugation at 6,000X g for 30 min, washed with PBS containing 1 mM PMSF using 30 mL/L of culture, and resuspended in column buffer (30 mL/L of culture), then frozen at -80 °C and thawed. 4. The following scheme summarizes purification from cells derived from 1 L of induced culture with all steps being performed at 4 °C. Steps 5 and 6 have been described previously (1,16), and step 7 is an unpublished method. 5. Resuspended cells are fractured in a French press and centrifuged at 27,000X g for 2 h (see Note 2). 6. The cleared supernatant is loaded onto a 12.5 mL column of Ni2+ -charged IMAC agarose that has been equilibrated with column buffer, washed with two volumes of column buffer, and eluted with a linear 0–200 mM imidazole gradient based on the same buffer but with only 10% glycerol (total gradient volume: 250 mL). Fractions are collected and assayed for pol III activity and the peak fractions pooled. 7. The IMAC pool is loaded onto a 10 mL MacroPrep High Q anion exchange column that has been equilibrated with MacroPrep column buffer, washed with three volumes of the same buffer, and eluted with a linear gradient of 0.1–0.6 M NaCl in the same column buffer (total gradient volume: 75 mL) (see Note 3). Fractions are collected and assayed for pol III activity, and the peak fractions pooled, aliquotted, and stored at -80 °C (see Note 4).

3.2. Preparation of DNA Substrates 3.2.1. Activated Calf Thymus DNA 1. Rinse two pairs of forceps with 90% ethanol, and use a flame to sterilize. Using the forceps, shred 800 mg of dry calf thymus DNA into small pieces and place into

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45000 40000

Counts

35000 30000

0.075 ug/mL

25000

0.15 ug/mL

20000

0.3 ug/mL

15000

0.6 ug/mL

10000 5000 0 0.00

5.00

10.00

15.00

Minutes

Fig. 1. Pilot-scale preparation of activated calf thymus DNA, described in Section 3.2.1.

2.

3. 4.

5.

a sterile flask. Add 400 mL sterile 50 mM Tris-HCl, pH 7.5 (to yield a 2 mg/mL solution), and shake at 125 rpm for 48 h at 4 °C. Perform a pilot run of DNAse digestion to determine the proper DNAse concentration and digestion time. Set up four glass tubes containing 2 mg/mL DNA, 5 mM MgCl2 , and DNAse at the following concentrations: 0.6, 0.3, 0.15, and 0.075 μg/mL. Incubate the tubes at 37 °C, remove 200 μL aliquots at 2.5, 5, 10, and 15 min, and inactivate DNAse by heating to 65 °C for 10 min and cooling samples on ice. Add 5 μL of each DNA sample to 15 μL DNA-free assay mix (see Section 2.2.1) and 5 μL pol IIIC at a 1:200 dilution in a 96-well assay plate and incubate for 10 min at 30 °C. Process samples as described in Section 3.3. Plot the results as counts vs. time for each DNAse concentration (e.g., see Fig. 1) and select the optimal DNAse concentration and digestion time. Scale up the reaction by placing 200 mL of the DNA solution into each of two 2 L flasks (see Note 5) and adding MgCl2 to a final concentration of 5 mM and DNAse to the concentration chosen in the pilot study (see Note 6). Incubate in a 37°C water bath with occasional swirling for the amount of time chosen in the pilot study, inactivate the DNAse by heating the flask in a 65°C water bath with constant swirling, and place flasks on ice to cool. Aliquot the activated DNA into 15 mL tubes and store at –20 °C for up to 10 years.

3.2.2. Synthetic Primer:Templates A number of combinations of synthetic template:primers can be used; however, we have found two combinations that produce good results with B. subtilis pol III: poly(dA):oligo(dT) and poly(dC):oligo(dG).

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In order to prepare these DNA substrates, a ratio of 5 A260 units of polymer [poly(dA) or poly(dC)] and 0.25 OD units of oligomer [dT12−18 or dG12−18 ] should be heated in the presence of 10 mM Tris-HCl, pH 7.6, at 50 °C for 10 min and allowed to cool to 25 °C over 10 min. 3.3. Assays Using Pol III 3.3.1. Standard Pol III Assay Using Activated DNA and Four dNTPs (One Labeled) Assays on both native and recombinant, purified B. subtilis pol IIIC are performed as previously described (13,14), with the exception of using a 96well plate format. 1. Each 25 μL assay contains 30 mM Tris, pH 7.5, 10 mM magnesium acetate, 4 mM dithiothreitol, 20% glycerol, with 25 μM dATP, dCTP, dGTP and 10 μM dTTP (3 H-labeled at 1.2 μCi/nmole) and 0.4 mg/mL activated calf thymus DNA as substrates in a polypropylene 96-well assay plate. 2. Assays are initiated by the addition of 0.025 to 0.06 units of pol III enzyme (1 unit is the amount required to incorporate 250 pmoles of [3 H]-dTMP in a standard assay), incubated for 10 min at 30 ºC, and terminated by the addition of 100 μL of cold 10% TCA, 10 mM sodium pyrophosphate (see Note 7). 3. Precipitated labeled DNA is collected on glass fiber filter plates that have been prewet with 50 μL ice-cold 1 M HCl, 100 mM sodium pyrophosphate. The plates are washed with 100 μL of the HCl solution followed by 100 μL of ice-cold 90% ethanol and then dried for 30 min under a heat lamp. 4. Liquid scintillation cocktail (30 μL OptiPhase Supermix) is added and the plate counted in a liquid scintillation counter (MicroBeta Trilux, Perkin Elmer) set to read the tritium signal for 15 s in a 96-well plate.

3.3.2. Assays Using Synthetic Primer:Templates Assay conditions using synthetic primer:templates are essentially the same as the assay described above for use with activated DNA. The differences are as follows: 1. A final template (polymer) concentration of 0.5 OD/mL is used in a 25 μL assay volume. 2. A single dNTP is present in each assay type: [3 H]-dTTP or [3 H]-dGTP (10 μM, 1.2 μCi/nmole) for use with poly(dA):oligo(dT) or poly(dC):oligo(dG), respectively (see Note 8). 3. The reaction must be incubated at 37 °C for 60 min to get optimal incorporation.

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3.3.3. Primer Extension Assays The primer extension assay is performed to assess the potential for incorporation into DNA of a dNTP mimic type of inhibitor and has been described in the literature (17–19). 1. Primer is labeled by incubating 0.125 A260 units with 500 μCi [32 P]-ATP (3000 Ci/mmol) and 5 units T4 polynucleotide kinase in 50 mM Tris-HCl, pH 7.6, 10 mM MgCl2 , 5 mM DTT, and 1 mM spermidine in a total volume of 50 μL for 60 min at 35 °C, followed by boiling for 2 min and desalting on a 250 μL Sephadex G-25 column equilibrated with 50 mM Tris-HCl, pH 7.6. 2. The desalted primer is annealed with an equal amount of template in labeling buffer minus spermidine, and the solution is heated to 50 °C for 5 min followed by slow cooling to room temperature over 60 min and quenching on ice. 3. Primer extension is performed by incubating 0.1–0.2 units of pol IIIC in the presence or absence of dGTP or a dGTP-like inhibitor at 35 °C for 30 min in 12.5 μL of a solution containing 20 mM potassium phosphate, pH 7.5, 8 mM MgCl2 , 4 mM DTT, 0.1 mM EDTA, and 50 μg/mL of labeled template:primer. The reaction is terminated by adding 4 μL of stop buffer, boiling for 2 min, and quenching on ice. 4. To visualize primer extension, a 5 μL sample is applied to a 12% polyacrylamide DNA sequencing gel and electrophoresed for 4 h at 1500 volts at constant power. The gel is then dried and exposed to a 14 × 17-in. sheet of Fuji X-ray film for 1–6 h and developed. 5. A sequencing ladder is run alongside the samples to identify the length of the primer. To create the ladder, 0.25 units of calf thymus terminal deoxynucleotidyl transferase is incubated with 1 μg of labeled primer in the presence of 10 μM BuPdGTP (20) or another dGTP-like inhibitor that is known to be incorporated.

4. Notes 1. The cloning and expression of B. subtilis polC can be performed using a number of different kits/protocols that are now available from companies such as Novagen (i.e., pET vectors). 2. Lysis of E. coli cells can also be achieved by sonication, although use of the French press is preferable. For example, we have used a Virsonic 475 Ultrasonic Cell Disrupter (Virtis Company) with a ½-in. probe, setting 5 (of a maximum of 10), using 3 × 10 s pulses to lyse cells. Always immerse the tube containing cells in a beaker of ice, and avoid foaming of the suspension to prevent protein denaturation. 3. A Mono-Q anion exchange column with an FPLC system (Pharmacia) has been used as the second column purification step in published versions of the pol III purification, but we have found the MacroPrep column to give an equivalent purification without requiring the purchase of an FPLC system. 4. Pol III is stably stored at -80 °C for many years but is susceptible to loss of activity upon multiple freeze/thaw cycles. The best way to store this enzyme is, after the

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8.

M. M. Butler and G. E. Wright initial storage in a large volume, e.g., 1 mL, upon thawing a tube for the first time, aliquot that volume into smaller units, e.g., 50 μL, for more frequent use. Perform DNAse treatment using a large flask and volume:liquid ratio to obtain the fastest possible heat inactivation. If the DNAse treatment lasts too long, activity begins to decrease rapidly. An example of a typical DNAse treatment pilot run is illustrated in Fig. 1. The scale-up conditions that were chosen in this particular experiment were to treat with 0.15 μg/mL of DNAse for 10 min. The standard 4-dNTP assay should be used to analyze inhibitors of an unknown mechanism. However, to identify inhibitors that compete with any of the dNTPs, a truncated assay should be used. This assay type requires the omission of one dNTP at a time, the use of activated DNA, and the inclusion of the [3 H]-labeled dNTP that is not the dNTP that has been omitted. This so-called truncated assay has been used to assay the anilinouracils, pol IIIC inhibitors whose activities compete with dGTP (7). Further characterization of inhibitors with a competitive mechanism of action (MOA) may be performed using synthetic primer:templates, such as in the assay described in Section 3.3.2. Detailed analysis of the MOA of pol IIIC inhibitors, i.e., the 6-(phenylhydrazino)uracils, has been accomplished using synthetic DNAs (9,21).

References 1 Kornberg, A., and Baker, T. (1992) DNA Replication, 2nd ed. W.H. Freeman and 1. Co., New York. 2 Wright, G. E., and Brown, N. C. (1999) DNA polymerase III: A new target for 2. antibiotic development. Curr. Opin. Anti-Infective Investig. Drugs 1, 45–48. 3 Tarantino, P. M., Zhi, C. G., Wright, E., and Brown, N. C. (1999) Inhibitors of DNA 3. polymerase III as novel antimicrobial agents against Gram-positive eubacteria. Antimicrob. Agents Chemother. 43, 1982–1987. 4 Ali, A., Aster, S. D., Graham, D. W., Patel, G. F., Taylor, G. E., Tolman, R. L., 4. Painter, R. E., Silver, L. L., Young, K., Ellsworth, K., Geissler, W., and Harris, G. S. (2001) Design and synthesis of novel antibacterial agents with inhibitory activity against DNA polymerase III. Bioorg. Med. Chem. Lett. 11, 2185–2188. 5 Yang, F., Dicker, I. B., Kurilla, M. G., and Pompliano, D. L. (2002) PolC-type 5. polymerase III of Streptococcus pyogenes and its use in screening for chemical inhibitors. Anal. Biochem. 304, 110–116. 6 Dervyn, E., Suski, C., Daniel, R., Bruand, C., Chapuis, J., Errington, J., Janniere, L., 6. and Ehrlich, S. D. (2001) Two essential DNA polymerases at the bacterial replication fork. Science 294, 1716–1719. 7 Wright, G. E., and Brown, N.C. (1976) Inhibition of Bacillus subtilis DNA 7. polymerase III by arylhydrazinopyrimidines. Novel properties of 2-thiouracil derivatives. Biochim. Biophys. Acta 432, 37–48.

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8 Alberts, B. M., Amodio, F. J., Jenkins, M., Gutmann, E. D., and 8. Ferris, F. L. (1968) Studies with DNA-cellulose chromatography. I. DNA-binding proteins from Escherichia coli. Cold Spring Harb. Symp. Quant. Biol. 33, 289–305. 9 Clements, J. E., D’Ambrosio, J., and Brown, N. C. (1975) Inhibition of Bacillus 9. subtilis deoxyribonucleic acid polymerase III by phenylhydrazinopyrimidines. Demonstration of a drug-induced deoxyribonucleic acid-enzyme complex. J. Biol. Chem. 250, 522–526. 10 Mackenzie, J. M., Neville, M. M., Wright, G. E., and Brown, N. C. (1973) Hydrox10. yphenylazopyrimidines: Characterization of the active forms and their inhibitory action on a DNA polymerase from Bacillus subtilis. Proc. Natl. Acad. Sci. USA 70, 512–516. 11 Lowry, O. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J. (1951) Protein 11. measurement with the Folin phenol reagent. J. Biol. Chem. 193, 265–275. 12 Ghosh, S., and Lowenstein, J. M. (1996) A multifunctional vector system for 12. heterologous expression of proteins in Escherichia coli. Expression of native and hexahistidyl fusion proteins, rapid purification of the fusion proteins, and removal of fusion peptide by Kex2 protease. Gene 176, 249–255. 13 Barnes, M. H., Leo, C. J., and Brown, N. C. (1998) DNA polymerase III of Gram13. positive eubacteria is a zinc metalloprotein conserving an essential finger-like domain. Biochem. 37, 15254–15260. 14 Barnes, M. H., and Brown., N. C. (1995) Purification of DNA polymerase III of 14. Gram-positive bacteria. Methods Enzymol. 262, 35–42. 15 Hammond, R. A., and Brown, N. C. (1992) Overproduction and purification of 15. Bacillus subtilis DNA polymerase III. Protein Expr. Purif. 3, 65–70. 16 Foster, K. A., Barnes, M. H., Stephenson, R. O., Butler, M. M., Skow, D. J., 16. LaMarr, W. A., and Brown, N. C. (2003) DNA polymerase III of Enterococcus faecalis: Expression and characterization of recombinant enzymes encoded by the polC and dnaE genes. Protein Expr. Purif. 27, 90–97. 17 Butler, M. M., Dudycz, L. W., Khan, N. N., Wright, G. E., and Brown, N. C. 17. (1990) Development of novel inhibitor probes of DNA polymerase III based on dGTP analogs of the HPUra type: Base, nucleoside and nucleotide derivatives of N2-(3,4-dichlorobenzyl)guanine. Nucleic Acids Res. 18, 7381–7387. 18 Khan, N. N., Wright, G. E., and Brown, N. C. (1991) The molecular mechanism 18. of inhibition of alpha-type DNA polymerases by N2-(butylphenyl)dGTP and 2(butylanilino)dATP: Variation in susceptibility to polymerization. Nucleic Acids Res. 19, 1627–1632. 19 19. Townsend, A. J., and Cheng, Y. C. (1987) Sequence-specific effects of ara-5-azaCTP and ara-CTP on DNA synthesis by purified human DNA polymerases in vitro: Visualization of chain elongation on a defined template. Mol. Pharmacol. 32, 330–339.

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20 Wright, G. E., and Dudycz, L. W. (1984) Synthesis and characterization of N2-(p20. n-butylphenyl)-2’-deoxyguanosine and its 5’-triphosphate and their inhibition of HeLa DNA polymerase alpha. J. Med. Chem. 27, 175–181. 21 Gass, K. B., Low, R. L., and Cozzarelli, N. R. (1973) Inhibition of a DNA 21. polymerase from Bacillus subtilis by hydroxyphenylazopyrimidines. Proc. Natl. Acad. Sci. USA 70, 103–107.

4 Methods to Identify and Characterize Inhibitors of Bacterial RNA Polymerase A. Simon Lynch and Qun Du

Summary RNA polymerase is essential to the viability of bacteria in all phases of growth and development and is a proven chemotherapeutic target as the cellular target of the rifamycin class of antibiotics. However, despite the characterization of multiple different classes of natural products that selectively target bacterial RNA polymerase, and the identification of a limited number of synthetic compound inhibitors, only agents of the rifamycin class have been developed and approved for human clinical use as antibiotics. Herein we describe a scintillation proximity assay (SPA) for identifying and characterizing inhibitors of bacterial RNA polymerases and that is applicable to de novo drug discovery programs through application of automated high-throughput screening methods. In addition, we describe gel electrophoresis-based methods that are applicable to the detailed characterization of inhibitors of transcriptional initiation or elongation by bacterial RNA polymerases.

Key Words: RNA polymerase; transcription; rifampin; Sigma factor; inhibitor; antibiotic; high-throughput screen.

1. Introduction Bacteria possess a single RNA polymerase (RNAP) enzyme that is responsible for the synthesis of all structural RNA (tRNA and rRNA) and messenger RNA (mRNA) species. RNAP is a complex macromolecular machine comprised of a multi-subunit catalytic “core” enzyme (2 ′ ) of ∼400 kDa that combines with an additional subunit, Sigma (), to form a “holoenzyme” capable of site-specific transcriptional initiation at cognate promoter elements (1). RNAP is essential for the propagation and maintenance of bacteria From: Methods in Molecular Medicine, Vol. 142: New Antibiotic Targets Edited by: W. Scott Champney © Humana Press Inc., Totowa, NJ

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in multiple different phases of growth and development, including disease pathogenesis and the adaptation to environmental stresses. Regulation of the levels or activity of alternate  factors, and other classes of transcriptional regulatory proteins, governs the temporal modulation of the transcriptome produced by RNAP and therein affects the gene expression changes necessary for transitions between different growth states. The essentiality of RNA polymerase in the growth and propagation of bacteria is reflected in the number of antibiotic molecules that have been identified that selectively target the enzyme. These include bacteriophageencoded proteins that act as anti-Sigma factors, exported polypeptide antibiotics like microcin J25 (2,3), and a structurally diverse group of secondary metabolite compounds including the rifamycins (4), myxopyronins (5), tiacumicins (6), streptolydigin (7), sorangicin (8), and GE23077 (9) identified from natural product sources in cell-based antimicrobial screens. To date, however, only members of the rifamycin class of antibiotics have been developed and approved for clinical use in humans. Biochemical high-throughput screening (HTS) methods have more recently been applied toward the identification of specific mechanistic classes of inhibitors of bacterial RNAP enzymes. Such efforts have yielded three structurally distinct classes of synthetic, small-molecule inhibitors that perturb RNAP function via effects on either holoenzyme assembly (10), initiation (11), or elongation (12,13). However, it remains to be determined whether these (or other) novel classes of RNAP inhibitors will yield antimicrobial agents of clinical utility. Herein we report methods that have been successfully employed in the identification and characterization of novel structural classes of inhibitors of the RNA polymerases of Escherichia coli and Staphylococcus aureus. 2. Materials 2.1. Bacterial Strains and Growth Media 1. S. aureus CB0842 is a derivative of strain RN4220 that bears an rpoC::His8 allele linked to a tetracycline-resistant determinant and was engineered by standard methods (A. S. Lynch, unpublished). 2. E. coli CU0311 is a derivative of BL21 (DE3)/pLysS (Novagen, EMD Biosciences) bearing a plasmid (pET24- A ) that expresses the S. aureus  A (PlaC, RpoD) protein in an isopropyl -D-1-thiogalactopyranoside (IPTG) inducible fashion (A. S. Lynch, unpublished). 3. Cation-adjusted Mueller Hinton (MH II) broth and agar medium. 4. Terrific Broth (TB) medium. 5. Luria Bertani (LB) medium.

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2.2. Protein Purification 1. Lysis buffer: 50 mM Tris-HCl (pH 7.9), 10 mM MgCl2 , 300 mM NaCl, 5% glycerol. See Note 1 regarding the preparation of buffers and solutions. 2. Complete Protease Inhibitor Tablets—EDTA-free (Roche Applied Science) 3. Lysostaphin. 4. 5 mL HiTrap HPLC Chelating Column (Amersham product # 17-0409-01 from GE Healthcare, Life Sciences). 5. Buffer A: 80 mM Tris-HCl, pH 7.9, 300 mM NaCl, 10% glycerol, 0.2 mM PMSF. 6. Buffer B: 80 mM Tris-HCl, pH 7.9, 300 mM NaCl, 10% glycerol, 0.2 mM PMSF, and 100 mM Imidazole. 7. RNAP storage buffer: 50 mM Tris-HCl, pH 7.9, 0.1 mM EDTA, 0.2 mM PMSF, 1.0 mM DTT, 50% glycerol. 8. TGED buffer: 50 mM Tris-HCl, pH 7.9, 5% glycerol, 0.1 mM EDTA, 0.5 mM DTT.

2.3. Scintillation Proximity Assays for Identifying RNAP Inhibitors 1. Yyttrium Silicate (YSi) RNA binding SPA beads (Amersham product # RPNQ0013, GE Healthcare, Life Sciences); see Note 2 for details regarding the storage and use of YSi SPA beads. 2. 96-well flat bottom, opaque, white polystyrene plates (Corning, VWR Cat # 29444-016); equivalent 384- or 1536-well Corning plates are recommended for screening in higher-density formats using automated high-throughput screening (HTS) platforms. 3. RNAP SPA buffer (5X): 200 mM Tris-HCl, pH 7.9, 500 mM KCl, 0.4% (v/v) PEG-8000, 50 mM MgCl2 , 25 mM DTT, 12.5% (v/v) glycerol. 4. Purified plasmid DNA template: pGL2B-T7A1 (see Fig. 2 and Note 3). 5. Ribonucleotide (NTP) mix (10X): 1 mM ATP, 1 mM CTP, 1 mM GTP, and 1 μM UTP. Prepared from 100 mM stocks (Promega), aliquotted, and stored at –20 ºC. 6. [5,6-3 H] Uridine 5’-triphosphate, ammonium salt; specific activity 1.1-2.2 TBq/mmol, 30–60 Ci/mmol (Amersham Radiochemicals product # TRK412, GE Healthcare, Life Sciences). 7. Nuclease-free water. 8. Dimethyl sulfoxide—99.9% ACS grade. 9. E. coli RNA Polymerase  70 -Saturated Holoenzyme (Product # S90250, Epicentre Biotechnologies). See Note 4. 10. S. aureus RNA Polymerase  A -Saturated Holoenzyme(prepared fresh as described in Section 3.1). 11. Test compounds and rifampin: Stocks should be prepared in 100% dimethylsulfoxide (DMSO), aliquotted, and stored at –20 ºC.

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A. S. Lynch and Q. Du (kDa) MW

EC

PA

SA

SP

MW

203

203

120

120

90

90

52

52

20

20

4

4

Fig. 1. SDS-PAGE analysis of recombinant RNAP enzymes. RNAP enzymes were purified from strains engineered to express RpoC (’) subunits that bear a carboxylterminal oligo-histidine tag (RpoC-His6−10 ) by immobilized metal affinity chromatography (IMAC): E. coli (EC), P. aeruginosa (PA), S. aureus (SA), and S. pneumoniae (SP). For each preparation, the two large catalytic subunits of the RNAP enzyme (RpoB and RpoC) are readily apparent. The molecular weight (MW) markers employed have the indicated sizes in kiloDaltons (kDa).

2.4. Gel-Based Assays for Characterizing RNAP Inhibitors 1. [-32P]-Cytidine 5’-triphosphate with specific activity of 10 μCi/μl (product # BLU508X from PerkinElmer Life and Analytical Sciences). 2. Polyacrylamide gel reagents: SequaGel Concentrate (product # EC-830), Diluent (EC-840) and Buffer (EC-835) from National Diagnostics Inc. 3. Recombinant RNasin Ribonuclease Inhibitor (product # N2515 from Promega Corp.). 4. Gel Loading Buffer II for Denaturing PAGE (Ambion product # 8547, Applied Biosystems). 5. Purified plasmid DNA template: pGL2B-T7A1 (see Fig. 2 and Note 3). 6. RNAP gel assay buffer (5X): 200 mM Tris-HCl, pH 7.9, 500 mM KCl, 0.4% (v/v) PEG-8000, 50 mM MgCl2 , 25 mM DTT, 12.5% (v/v) glycerol. 7. S. aureus gel assay ribonucleotide (NTP) mix (10X): 6 mM ATP, 6 mM UTP, 0.01 mM CTP. Prepared from 100 mM stocks (Promega), aliquotted, and stored at –20 ºC. 8. E. coli gel assay ribonucleotide (NTP) mix (10X): 6 mM ATP, 6 mM UTP, 0.001 mM CTP. Prepared from 100 mM stocks (Promega), aliquotted, and stored at –20 ºC.

3. Methods The identification and characterization of RNAP inhibitors require access to highly purified enzyme preparations that exhibit high-specific activity in vitro. Despite the availability of E. coli RNAP from a number of commercial sources,

RNA Polymerase assays

41 -35

-10

+1

PT7A1 Psea

GAGTATTGACTTAAAGTCTAACCTATAGGATACTTACAGCCATA TATTATAGACAAGTATAAAAAAGGTATAGTAATATATGTATATA

PTAC18

CGCTATTGACATTATAGTGGTACCGCTTATATAATCTCAATTAT

Bgl II

Hind III

Promoter

G-less Nco I

BamH I

Luciferase Sal I

Fig. 2. Diagrammatic representations of transcription templates used for the identification and characterization of bacterial RNAP inhibitors. The promoter elements shown were combined by PCR methods with a 272 nucleotide G-less cassette element derived from the vector p(C2AT)19 (24) and cloned between the Bgl II and Hind III sites of the pGL2-Basic plasmid (Promega Corp.; Genbank Accession Number X65323). In pGL2B-T7A1-Gless, the bacteriophage T7 derived PT 7A1 promoter is utilized by RNAP enzymes from both Gram-positive and Gram-negative species. In pGL2B-sea-Gless, the Psea promoter is derived from the transcriptional regulatory region of the Staphylococcus enterotoxin-A (sea) gene promoter and is efficiently utilized by S. aureus  A RNAP enzyme. In pGL2B-TAC18-Gless, the PTAC18 promoter is a synthetic trplacUV5 element that is efficiently utilized by E. coli  70 RNAP enzyme and other Gram-negative RNAP enzymes (A. S. Lynch et al., unpublished). Using negatively supercoiled DNA as a substrate, and optimal transcription conditions, initiation from the promoter elements in the presence of ATP, CTP, and UTP (but no GTP) yields a single RNA species that can be readily separated by gel electrophoresis and quantified. Alternately, linear DNA substrates may be generated by restriction enzyme cleavage and transcription undertaken in the presence of GTP.

only the  70 -saturated holoenzyme preparation from Epicentre (Madison, WI) has proven suitable for detailed mechanistic studies. However, to facilitate the relatively inexpensive purification of large quantities of high-specific-activity RNAP enzymes for use in HTS, we have adopted a strategy wherein recombinant strains of target bacteria are engineered that express an RpoC (’) subunit that bears a carboxyl-terminal oligo-histidine tag (RpoC-His6−10 ). Purification of oligo-histidine tagged core RNAP preparations via immobilized metal affinity chromatography (IMAC) followed by reconstitution with recombinant  factor yields RNAP holoenzyme forms that exhibit high-specific activity in vitro. Similar methods have been reported for Streptococcus pneumoniae (14), Bacillus subtilis (15,16), Streptomyces coelicolor and S. lividans (6), and have also proven applicable to studies of RNA polymerases from Pseudomonas aeruginosa, Escherichia coli, Haemophilus influenzae, and Mycobacterium smegmatis (A. S Lynch et al., unpublished). Herein we describe in detail the purification and reconstitution of S. aureus  A RNAP holoenzyme by

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methods that are readily adaptable for the preparation of similar recombinant enzymes from other bacteria. A number of biochemical HTS assay methods have been reported that are broadly applicable to the identification of novel inhibitors of RNAP enzymes including both (1) non-homogenous methods involving RNA product capture (17,18) and (2) homogenous methods that utilize either the scintillation proximity assay (SPA) (19,20) or fluorescence-based molecular beacon (21,22) technologies for RNA product detection. Herein we describe in detail the use of a 96-well SPA-based HTS assay employing either commercially supplied E. coli  70 or purified S. aureus  A RNAP holoenzymes that can be run manually or in an automated fashion. This assay is readily adaptable for implementation in 384- or 1536-well automated formats. A variety of assays have been reported that are applicable to the detailed characterization of different mechanistic classes of inhibitors of bacterial RNAP enzymes (23). Herein we describe methods for the characterization of inhibitors of transcriptional initiation or elongation and that are based on direct quantification of either a single (or multiple) radiolabeled RNA product(s) following resolution by gel electrophoresis. The transcription template employed in the assays described in detail bears the bacteriophage T7A1 promoter, which is efficiently utilized by bacterial RNAP holoenzymes of both Gram-negative and Gram-positive origin; however, alternate transcription templates can be readily substituted that are applicable to studies of specific RNAP enzymes. Hence, while the methods described are focused on characterization of inhibitors of either E. coli  70 or S. aureus  A RNAP holoenzymes, they are readily adaptable for similar studies of RNAP enzymes from other bacterial sources. Finally, DNA templates that incorporate a “G-less” cassette element (24) allow for detailed kinetic studies of RNAP inhibitors through analysis of the yield of synthesis of a unique RNA product formed by a single round of transcription. 3.1. Purification of Recombinant RNA Polymerase 1. A fresh culture of S. aureus CB0842 (rpoC::His8 ::TetR ) is prepared from frozen glycerol stocks by recovery onto an MHII agar plate (containing 2.5 μg/mL tetracycline) and incubation at 37 ºC overnight. 2. 400 mL of Terrific Broth (TB) medium containing 1 μg/mL tetracycline is inoculated with a single colony of CB0842, and the culture is grown overnight at 37 ºC with aeration. 3. Aliquots of the fresh overnight culture of CB0842 are used to inoculate 8 L of TB at a starting OD600 of ˜0.1, and the cultures are grown at 37 ºC with aeration for 6–8 hours until mid-exponential phase (corresponding to an OD600 of 3 to 4).

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4. The bacterial cells are then harvested by centrifugation at 5000 RPM (4000X g) for 20 min in a Sorvall H6000A rotor. This step and all subsequent steps are carried out at 4 ºC or on ice unless stated otherwise. 5. Cell pellets are resuspended, washed in Lysis buffer, and consolidated into a single pellet by centrifugation at 12,000 RPM (12,000X g) for 20 min in a Sorvall SLA-1500 rotor. The cell pellet is then either immediately processed as described below or flash frozen in liquid nitrogen and stored at –80 ºC. 6. If previously frozen, the cell pellet (typically ∼50 g) is gradually thawed and suspended in 170 mL of Lysis buffer. Four complete protease inhibitor tablets (EDTA free) and 20 mg of lysostaphin (as a dry powder) are then added, and the bacterial cell suspension is incubated at 37 ºC for 1 h, with occasional mixing, and then placed on ice. 7. The viscosity of the bacterial cell solution is then reduced by sonication: 5–10 cycles for 30 s on the maximum power setting with a Branson Digital Sonifier (or equivalent) with intermittent cooling on ice. Cell lysis is then completed by 4–6 cycles of treatment with a microfluidizer (Microfuidics Corp.) at 80–100 pounds per square inch (psi) of pressure at 4 ºC. 8. Insoluble cell debris is then removed by centrifugation at 12,000 RPM (12,000X g) for 20 min in a Sorvall SLA-1500 rotor. 9. RNAP enzyme is purified by IMAC using two 5 mL HiTrap HP columns (in series) that are charged with CoCl2 and equilibrated in lysis buffer. The supernatant from Step 8 is loaded onto the HiTrap HP columns using an ÄKTA FPLC system (Amersham, or equivalent) and washed with 20 column volumes of Buffer A, and the protein is then eluted with a 20 column volume linear 0–100 mM imidazole gradient that is produced by mixing Buffer A and Buffer B. 10. Peak column fractions are identified by SDS-PAGE analysis, pooled, and dialyzed against RNAP storage buffer containing 0.2 mM PMSF. 11. Following dialysis, the concentration of the purified protein preparation is determined and aliquots flash frozen in liquid N2 and stored at –80 ºC. The final protein yield is typically 20 mg; see Fig. 1 for SDS-PAGE analysis of a representative preparation.

3.2. Purification of Recombinant Sigma Factor 1. A fresh culture of E. coli CU0311 is prepared from frozen glycerol stocks by recovery onto an LB agar plate containing kanamycin (50 μg/mL) and chloramphenicol (35 μg/mL) and by incubation at 37 ºC overnight. 2. 400 mL of TB medium containing kanamycin (50 μg/mL) and chloramphenicol (35 μg/mL) is inoculated and grown overnight at 37 ºC with aeration. 3. 1 L of antibiotic supplemented TB medium is then inoculated with the overnight culture at a starting OD600 of 0.05, and the culture is grown at 37 ºC with aeration to an OD600 of 0.4. Expression of the recombinant S. aureus  A (RpoD) protein is then induced by the addition of IPTG to 0.5 mM.

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4. After 3 h at 37 ºC with vigorous aeration, the bacterial cells are harvested by centrifugation for 10 min at 5000 RPM (5000X g) at 4 ºC. 5. Recovered cells are then washed in 50 mL of cold TGED buffer containing 200 mM NaCl and pelleted by centrifugation at 5000 RPM (5,000X g) at 4 ºC, and the resulting cell pellet weighed and then flash frozen in liquid N2 and stored at –80 ºC. A typical yield is 4 g of cells (wet weight) per liter of induced cell culture. 6. The cell pellet is slowly thawed and suspended in 16 mL of cold TGED buffer containing 200 mM NaCl per 4 g of cells, and the following are then added: dithiothreitol (DTT) to 1 mM, PMSF to 0.2 mM, and one complete (-EDTA) protease inhibitor cocktail tablet (per 50 mL of volume). 7. The cells are lysed by incubation on ice for 30 min followed by sonication. The majority of the S. aureus  A protein is present in inclusion bodies, which are harvested by centrifugation of the lysate at 12,000 RPM (12,000X g) at 4 ºC for 30 min in a Sorvall SS-34 rotor. 8. The resulting inclusion body pellet is suspended in 20 mL TGED buffer containing 6 M guanidine-HCl and the solution mixed for 30 min at 4 ºC. Insoluble material is then removed by centrifugation for 20 min at 4 ºC at 12,000 RPM (12000X g) in a Sorvall SS-34 rotor. 9. To re-nature the  A protein, the supernatant is dialyzed extensively against TGED buffer containing 100 mM NaCl, 0.5 mM DTT, and 0.2 mM PMSF. 10. Following dialysis, insoluble material is removed by centrifugation at 12,000 RPM (17,000X g) in a Sorvall SS-34 rotor for 30 min at 4 ºC. The re-natured  A protein is then precipitated with ammonium sulfate, and the protein pellet recovered by centrifugation at 14,000 RPM (23,000X g) in an SS-34 rotor. The protein pellet is suspended in 1–2 mL of TGED buffer containing 1 mM DTT and 0.2 mM PMSF and further purified by size-exclusion chromatography on a Sephacryl S-200 HR FPLC column (Amersham) equilibrated with TGED buffer containing 200 mM NaCl and 0.2 mM PMSF. 11. Column fractions containing peak fractions of the  A protein are identified by SDS-PAGE analysis, pooled, and dialyzed against TGED buffer containing 100 mM NaCl and 50% glycerol. 12. Following dialysis, the concentration of the purified protein preparation is determined and aliquots flash frozen in liquid N2 and stored at –80 ºC. The final protein yield is typically 30 mg per liter of induced cells.

3.3. Reconstitution of RNA Polymerase Holoenzyme 1. Aliquots of the purified recombinant S. aureus RNAP enzyme and  A protein are thawed on ice from –80 ºC stocks and then combined at the time of use in transcription reactions. In standard assays,  A protein is included in a 5-fold molar (stoichiometric) excess over the core RNAP enzyme to yield a “Sigma-saturated”

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holoenzyme form that exhibits high specific activity in vitro. See Note 5 for details of methods used to estimate the relative stoichiometry of RNAP subunits. 2. If reconstitution of an alternate S. aureus RNAP holoenzyme form is desired, the recombinant RNAP enzyme must be further purified by repeated passage through a phosphocellulose column to yield a core RNAP enzyme preparation devoid of contaminating Sigma factors (see Note 6). 3. Prior to use in mechanistic studies of RNAP inhibitors, reconstituted S. aureus RNAP holoenzymes should be characterized to ensure that they exhibit site-specific initiation of transcription from DNA templates bearing a cognate S. aureus promoter element (see Section 3.5).

3.4. High-Throughput Method to Identify RNA Polymerase Inhibitors 1. Prepare a test compound plate at 5X concentration in a clear 96-well polypropylene plate with rifampin included in a 6- to 12-well titration covering the final concentration range of 0.25–250 ng/mL. 2. On ice, prepare mix 1 with 25 μL per assay well: 10 μL of RNAP SPA buffer (5X), 100–300 ng of S. aureus  A -saturated RNAP holoenzyme and nuclease-free water (to 25 μL). 3. On ice, prepare mix 2 with 15 μL per assay well: 5 μL NTP stock (10X), 250 ng pGL2B-sea DNA, and 0.2 μL [3 H] UTP, and nuclease-free water (to 15 μL). Use of a plastic assay trough is recommended if more than 20 assay wells are to be used. 4. Aliquot 25 μL of mix 1 to appropriate test assay wells of a 96-well flat bottom, white polystyrene plate. Use a multi-channel pipette if pipetting from an enzyme trough. 5. Add 10 μL of each test compound (pre-diluted as in 1) to the appropriate test wells and then 15 μL of mix 2 to each assay well. Gently shake the plate by hand and incubate at 25 ºC for 40 min (see Note 7). 6. Freshly prepare SPA bead binding buffer with 100 μL per assay well: 99.5 μL 166 mM NaCitrate (pH 2.4) plus 0.5 μL YSi RNA binding SPA beads (suspended by vortex immediately before use). 7. Add 100 μL of SPA bead mixture (from step 6) to each well and seal the plate with a clear adhesive seal (Packard). Allow the plates to stand undisturbed for at least 30 min (to allow the beads to settle) prior to reading the plates (as below). Note that plates are stable at room temperature for at least 24 h. 8. Quantify the RNA synthesis products by reading the plate in a microplate scintillation counter (see Note 8). Figure 3a shows enzyme inhibition curves obtained for rifampin with either E. coli  70 or S. aureus  A RNAP holoenzymes. Note that 50% inhibition concentration (IC50 ) values determined for rifampin in this assay for these enzymes typically fall in the 5–20 nM range and, as such, are consistent with literature precedents and data from gel-based assays (see below).

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A. S. Lynch and Q. Du SPA Assay Data 120

S. aureus σA Rif-S

% Remaining Activity

100

E. coli σ70 RpoC::His6

80 60 40 20 0 0.1

1

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Rifampin Concentration (Log nM)

Gel Assay Data

% Remaining Activity

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E. coli σ70 (Epicentre)

40

E. coli σ70 RpoC::His6

20 0 0.1

1

10

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Rifampin Concentration (Log nM)

Fig. 3. Analysis of data from the SPA and gel format assays. Figure 3a shows titrations of rifampin in the SPA format assay obtained for reconstituted forms of the RpoC::oligo-histidine tagged variants of the S. aureus  A and E. coli  70 RNA polymerase holoenzymes. Analysis of the data using Prism version 3.03 (GraphPad Software Inc.) yielded apparent 50% inhibition concentration (IC50 ) values of 6 and 9 nM for the S. aureus and E. coli enzymes, respectively. Figure 3b shows titrations of rifampin in the gel format assay obtained for the indicated RNA polymerase holoenzymes using the pGL2B-T7A1-Gless template in reactions conducted in the absence of GTP. Analysis of the data using Prism yielded apparent IC50 values of 5.5, 9.9, and 10.5 nM for the E. coli  70 (RpoC::His6 ), E. coli  70 (Epicentre), and S. aureus  A (rifampin-sensitive, RpoC::His8 ) enzymes, respectively. Also shown is a titration of rifampin against a rifampin-resistant variant of the S. aureus  A RpoC::His8 enzyme that

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3.5. Gel-Based Method to Characterize RNA Polymerase Inhibitors 1. Prepare a test compound plate at 10X concentration in a clear 96-well polypropylene plate (or 8-well PCR strip plates) with rifampin included in a 6- to 12-well titration covering the final concentration range of 0.25–250 ng/mL. 2. Transcription reactions are generally carried out in 96-well trays in a final volume of 50 μL. On ice, combine the following per 50 μL assay well: nuclease-free water (as necessary to achieve a volume of 50 μL), 10 μL of RNAP gel assay buffer (5X), 1.5 μg (E. coli) or 1.5 μg (S. aureus) of freshly re-constituted -saturated RNAP holoenzyme, 500 ng of pGL2B-T7A1 DNA, 2 μCi of [-32P]-CTP, and 5 μL of test compounds (or DMSO). The plate is then incubated at 37 °C for 5 min. See Note 9 regarding the order of addition of reagents, and see Note 10 regarding the choice of DNA template. 3. Transcription is then initiated by the addition of 5 μL of the appropriate 10X NTP mix (per 50 μL assay reaction) and the plate incubated at 37 °C for 15 min (E. coli) or 30 min (S. aureus). 4. Reactions are terminated by the addition of 50 μL of gel loading solution, and the reaction products maintained on ice until just prior to gel electrophoresis analysis. For long-term storage, reaction products are stored frozen at −20 °C. 5. Reaction products are heated in a 120 °C sand block for 2 min and then placed on ice. Five to 30 μL aliquots of each are then loaded and separated on preheated 4% polyacrylamide/urea/TBE denaturing PAGE gels. Dried electrophoresis gels are exposed to phosphorimager cassettes and analyzed by quantitative phosphorimaging. Figure 3b shows enzyme inhibition curves obtained for rifampin with various E. coli  70 and S. aureus  A RNAP holoenzyme preparations. Note that IC50 values determined for rifampin in this assay for these enzymes typically fall in the 5–20 nM range and, as such, are consistent with literature precedents.

4. Notes 1. Unless otherwise stated, all solutions and buffers should be prepared using highquality de-ionized water that is free ( 30S and 32S —> 45S —> 50S) at high temperature. Compounds inhibiting DnaK are therefore expected to hinder ribosome biogenesis at temperatures above 37 °C. Among a myriad of other functions (7), DnaK is also implicated in the assembly and disassembly of protein and nucleoprotein complexes, and it was therefore tempting to check whether DnaK also plays a role in the in vivo -complementation of -galactosidase. This is an example of protein fragment complementation in which two segments of a protein associate noncovalently to form a functional structure. This was indeed the case (8), and the observation of a role, either direct or indirect, for DnaK in -complementation provides an easy and original phenotype (measurement of - complemented -galactosidase activity) to detect functional changes in DnaK, and therefore to screen routinely for compounds altering its activity. Surprisingly, this target has been exploited very little until now, perhaps because of lack of an easy assay to monitor in vivo the biological activity of DnaK. This tool now exists. We reasoned that DnaK, which catalyzes specific inter- and intramolecular protein interactions, should therefore participate in the -complementation of -galactosidase, a paradigm for the formation of a quaternary structure. Indeed, we have shown, by comparison between a dnaK+ and a dnaK756-ts mutant of E. coli, that functional DnaK is necessary for an enzymatically active -complemented ß-galactosidase (8). This novel and easy phenotype (blue versus colorless bacterial colonies on X-Gal-containing LB plates) and a measure of ß-galactosidase activity in bacterial crude extracts therefore offer a reliable and routine tool to monitor the activity of DnaK in vivo and to screen for potential inhibitors. 2. Materials 2.1. E. coli Strains and Growth Media 1. Ribosome assembly defects are generally exacerbated in E. coli strains devoid of stringent control (relaxed phenotype), and therefore any dnaK+ , relA strain such as AL26 (relA251: :kanR ) (4) or MC4100(relA1) (5) is suitable for studying the pattern of labeled ribosomal particles by sedimentation analysis. 2. Strains harboring either a mutant dnaK allele (AL19 = AL26dnaK756-ts) (3) or a dnaK null allele (BB1553 = MC4100dnaK52: :cmR , sidB1 (5)) (see Note 1) can be used as control strains exhibiting typical ribosome assembly defects when

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bacteria are radiolabeled at the non-permissive temperature (43 °C for AL19 ; 38 °C or more for BB1553 (see Note 2)). 3. Both minimal medium A (10), supplemented with 0.2% glucose, 1μg/mL thiamine, and 0.2% casaminoacids, and MOPS medium (11) can be used for labeling bacteria with [ 3 H]-uridine in liquid cultures. 4. To assay DnaK-dependent - complementation of -galactosidase, any dnaK+ , lacZM15 strain such as JM83 (12), DH5, NM522, or TR14 (8) transformed by any plasmid encoding the LacZ-peptide such as pUC18/19 or pWSK129 (13) can be used. Strains harboring either a mutant dnaK allele (TR20 = TR14dnaK756-ts) (8) or a dnaK null allele (JM83 = JM83dnaK52: :cmR ) (8) transformed by any plasmid encoding the LacZ-peptide listed above ( pUC18, pWSK129) can be used as control strains. These are typically unable to sustain -complementation when grown at 30 °C (in the case of JM83 ; see Note 3), or at the semi-permissive temperature of 37 °C to 40 °C (in the case of TR20; see Note 4).

2.2. Labeling of Bacteria with [3 H]-Uridine The [ 5-3 H]-uridine from Amersham (TRK178, 1 mCi/mL) of high specific activity (26 Ci/mmol) is isotopically diluted with non-radioactive uridine (Sigma, U-3750), to be added to the culture at a final concentration of 3 μM and1 μCi/mL. Incubation of bacteria is continued for 1 h (see Note 5). 2.3. Sedimentation of Ribosomal Subunits and Precursor Particles by Sucrose Gradient Centrifugation 1. Alumina (Alcoa, A305) is used to grind bacteria, and the bacterial crude extract is prepared in TMNSH buffer: 10mM Tris-HCl, pH 7.4, 10 mM magnesium acetate, 60 mM ammonium chloride, 1 mM dithiothreitol. 2. A stock solution of 50% (w/w) sucrose is prepared by adding 1 kg sucrose to 1 L of de-ionized water and is autoclaved for 20 min at 110 °C (not more, to avoid carbonizing the sucrose). Linear sucrose gradients are made from 10% and 30% sucrose solutions prepared either in TMNSH buffer (for sedimentation of ribosomes in association conditions) or in TMNSH buffer containing 400 mM NaCl (for sedimentation of ribosomes in dissociation conditions). 3. Ultracentrifugation is performed in an SW28 rotor (Beckman), which can take six 35 mL sucrose gradients or in an SW41 rotor (Beckman) with six 11 mL sucrose gradients. 4. After centrifugation, sucrose gradients are collected in about 30 fractions by aspirating from the bottom of the gradient with a peristaltic pump. Optical density (A260 ) is measured first, and then the radioactivity contained in each fraction is measured by precipitating the whole fraction with 5% trichloracetic acid, collecting the acid-insoluble radioactivity on Millipore filters, and counting the [3 H] c.p.m. by liquid scintillation counting.

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2.4. -galactosidase Activity 1. LB plates (Luria-Bertani medium solidified with 1.5% agar) containing 100 μg/mL X-Gal, the appropriate antibiotic depending on the resident plasmid (100 μg/mL ampicillin or 25 μg/mL kanamycin), and IPTG at various concentrations (from 0 to 1 mM). 2. X-Gal at 20 mg/mL in dimethyl formamide. 3. Z buffer: 60mM Na2 HPO4 , 40mM NaH2 PO4 , 10mM KCl, 1mM MgSO4 , 50 mM -mercaptoethanol. 4. Toluene (or chloroform and 0.1% SDS). 5. Ortho-nitrophenyl--galactoside (ONPG), a 4 mg/mL solution in 100 mM potassium phosphate buffer, pH 7. 6. 1 M Na2 CO3 solution.

3. Methods 3.1. Analysis of the Ribosome Assembly Pattern 1. To label ribosomes synthesized by the strain under study (wild-type or mutant dnaK, with a resident plasmid or not), liquid cultures (50 mL to 200 mL) are inoculated with 0.01 volume of an overnight preculture grown in the same medium at 30 °C. 2. Incubation at 30 °C in a shaking water bath is continued for 3 to 5 h, and the A600 /mL of the culture is measured each hour. 3. For labeling at 30°C, [3 H]-uridine is added to the culture at an A600 /mL between 0.3 and 0.6, and incubation is continued for 1 h. 4. For labeling at a non-permissive temperature, the culture is transferred to a second water bath set up at the required temperature, and shaken for 1.5 h (see Note 6) before adding the labeled uridine as above. Incubation is then continued for 1 h. 5. 1 mL of bacterial samples are withdrawn and acid-insoluble radioactivity is measured, to estimate the amount of tritated uridine incorporated into RNA. Labeling at 43 °C generally results in 105 to 106 [3 H] c.p.m. incorporated in the culture, about 2–3-fold less than that obtained at 30 °C. 6. Aliquots of the culture and of the overnight culture are also streaked on plates to verify the expected phenotypes of the strain under study, like antibiotic resistance or thermosensitivity. 7. Bacteria are collected by centrifugation, washed with TMNSH buffer, and the bacterial pellets (usually 50 to 200 mg) are kept frozen at –20 °C in 15 mL Corex tubes. 8. Bacterial crude extracts are prepared by grinding bacteria in the Corex tubes with a glass rod in the presence of twice their weight of alumina. They are resuspended in 1 to 2 mL of TMNSH buffer, then centrifuged at 10,000 rpm (12,000X g) for 20 min in a Sorvall centrifuge at 3 °C to pellet the alumina, unbroken cells, and cell debris. The supernatants are immediately layered on sucrose gradients.

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3.2. Sucrose Gradient Centrifugation 1. Crude extracts (about 105 [3 H] c.p.m.) are mixed at 4 °C with 2 A260 units each of non-radioactive wild-type E. coli 50S and 30S ribosomal subunits and layered onto 11 mL 10% to 30% linear sucrose gradients prepared in TMNSH buffer. The gradients are centrifuged at 3 °C in a Beckman SW41 rotor for 16 h at 26,000 rpm (110,000X g) for analysis under ribosome association conditions. 2. For analysis under ribosome dissociation conditions, samples prepared as above are adjusted to 400 mM NaCl and layered onto 35 mL 10% to 30% linear sucrose gradients prepared in 400 mM NaCl-containing TMNSH buffer, and centrifuged at 3 °C in a Beckman SW28 rotor for 17 h at 27,000 rpm (110,000X g). Each gradient is then collected in about 30 fractions whose optical density (A260 /mL) and [3 H] radioactivity are measured, as described in Section 2. A typical result is shown in Fig. 1.

Fig. 1. Sedimentation profiles of ribosomes and their subunits prepared from strains AL26 (dnaK+ ) (A and B) , and AL19 (dnaK756-ts) (C and D) labeled with [3 H]-uridine at 43°C. Ionic conditions in sucrose gradients promote either association of ribosomal subunits (A and C) or their dissociation (B and D). Sedimentation is from right to left. A260 /mL, open circles. [3 H] c.p.m, closed circles. (Taken from (4), with kind permission of Springer Science and Business Media.)

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3.3. -galactosidase Activity 1. The -galactosidase activity can be estimated (from the blue vs. colorless phenotype of the bacterial colonies) by streaking strains on LB plates containing 100 μg/mL X-Gal, the appropriate antibiotic depending on the resident plasmid (100 μg/mL ampicillin or 25 μg/mL kanamycin), and IPTG at various concentrations (from 0 to 1 mM). 2. -galactosidase-specific activities of bacteria grown in liquid cultures (in LB medium) are measured and expressed in Miller units following standard procedures (10). -galactosidase activity can be measured after lysis of 1 mL samples of bacteria kept on ice by the addition of one drop of toluene or chloroform and 0.1% SDS. -galactosidase activity can also be measured the following day if desired. 3. Liquid cultures in LB medium or in minimal medium containing 0 to1 mM IPTG are inoculated at low cell density by a 1/200 or 1/500 dilution of an overnight preculture in the same medium. Bacterial growth is followed by measuring the A600 /mL, and several bacterial samples (four or five) are withdrawn between 0.25 (exponential growth) and 1.5 (stationary phase). The aliquots of the cultures are added to the assay medium, ice-cold Z buffer in 5 mL glass tubes to give a final volume of 1 mL. If high levels of -galactosidase are expected, add 0.1 mL of culture to 0.9 mL of Z buffer. In case of low levels, add 0.5 mL of culture to 0.5 mL of Z buffer. 4. Samples are kept on ice and bacteria are permeabilized by vortexing with one drop of toluene (or one drop of chloroform and one drop of 0.1% SDS). -galactosidase activity can be measured the following day; in this case, samples are kept overnight in the cold (to allow toluene to evaporate). 5. Tubes are placed in a water bath at 28 °C for 5 min, and the reaction is started (record the time) by adding 0.2 mL ortho-nitrophenyl--galactoside (ONPG). When sufficient yellow color has developed, the reaction is stopped by adding 0.5 mL of a 1 M Na2 CO3 solution (record the time again). 6. Transfer samples to Eppendorf tubes and spin down the cell debris in a small centrifuge for 2 min at maximal speed. 7. Transfer supernatants to 1 cm pathlength cuvettes and record the optical density at 420 nm. The A420 /mL should be ideally between 0.2 and 1.The -galactosidasespecific activity is expressed per A600 /mL unit of the bacterial culture, i.e., in Miller units (10), as follows = 1000 × A420 /mL/t × v × A600 /mL, where t is the time of the reaction in minutes, and v the volume of culture used in the assay in mL. In general, -galactosidase-specific activity should be constant during the exponential growth phase of the bacterial culture.

3.4. Screening for Antibacterial Compounds Targeting DnaK 1. An E. coli strain designed for a routine protocol (TR14, for example) is dnaK+ and harbors a chromosomal copy of the lacZM15 allele, which carries a partial deletion that does not disturb the open reading frame. This lacZM15 gene product (called the  fragment) lacks amino acid residues 11 through 41 and forms an inactive

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dimer (wild-type ß-galactosidase is a homotetramer). This strain is transformed by a plasmid encoding the 7500 dalton -fragment of the ß-galactosidase. In the presence of the -fragment, the - fragment reconstitutes a homotetramer with enzymatic activity. This activity is easily monitored with the two chromogenic substrates of the ß-galactosidase, X-Gal (producing blue colonies) and ONPG (hydrolyzed to the yellow-colored orthonitrophenol by bacterial crude extracts). 2. In a typical routine assay, the TR14 strain is grown at 37 °C in LB medium in the presence of the appropriate antibiotics (depending on the plasmid; see Note 7) and 100 μM IPTG, the lactose operon inducer (see Note 8). Compounds to be assayed are introduced at the final concentration of 100 μg/mL in a first attempt. When bacteria reach the stationary phase, cell growth (A600 /mL) and ß-galactosidase activity (A420 /mL of crude cell extracts after hydrolysis of ONPG) are measured. 3. The assay can be miniaturized to be installed on a Beckman Biomec 2000 robot allowing the use of 96-well plates for screening. Each set of experiments should include the control strain TR20pUC18 or TR20pWKS129 harboring the dnaK756-ts allele, which abolishes the -complementation of the ß-galactosidase at 37 °C (but not at 30 °C; see Fig. 2).

Fig. 2. -Complementation of -galactosidase in dnaK756 (Ts) and dnaK+ strains plated at 30 °C and 37 °C. Strains TR14 (dnaK+ ) pUC18 and TR20 [dnaK756(Ts)] pUC18 were streaked on two LB plates containing 100 μg of X-Gal per mL, 1 mM IPTG, and 100 μg of ampicillin per mL and incubated for 48 h at 30 °C or 37 °C. pUC18 is a high-copy-number plasmid with a ColE1 replication origin, expressing a LacZ -peptide and resistance to ampicillin. (Taken from (8), with permission of American Society for Microbiology.)

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4. Compounds with a positive signal can be studied over a wide range of concentrations, to determine their minimal inhibitory concentration (M.I.C.) for -galactosidase activity. They can also be checked to see whether a plasmid-driven overexpression of DnaK (for example, plasmid pKP31; see Fig. 3) reverses the inhibitory effects of these compounds. In order to confirm a specific inhibition of DnaK and not of the ß-galactosidase itself, positive compounds can be tested against an IPTG-induced wild-type lacZ+ strain where no inhibition is expected. Finally, they can be tested in other microbiological assays that necessitate a functional DnaK, but are unrelated to the -complementation of the ß-galactosidase, such as the lytic multiplication of bacteriophage , the shut-off of the heat-shock response (7), or the DnaK-dependent ribosome assembly. 5. An obvious limitation of the methodology described here is the selective permeability barrier of the outer membrane of E. coli toward many compounds (large peptides, for example). To circumvent this problem, the -complementation system

Fig. 3. DnaK-dependent -complementation of -galactosidase. Strains TR14 (dnaK+ ) [pWSK129 pBR322], TR14 (dnaK+ ) [pWSK129 pKP31], TR20 [dnaK756(Ts)][pWSK129 pBR322], and TR20 [dnaK756(Ts)][pWSK129 pKP31] were streaked on two LB plates containing X-Gal, IPTG, ampicillin, and 25 μg/mL of kanamycin, which were incubated for 48 h at 30 or 37 °C. pWSK129 is a lowcopy-number plasmid with a pSC101 replication origin, expressing a LacZ -peptide and resistance to kanamycin. Plasmid pKP31 (ColE1 replication origin, resistance to ampicillin), which overexpresses dnaK+ , is compatible with pWSK129. pBR322 is the empty vector corresponding to pKP31. (Taken from (8), with permission of American Society for Microbiology.)

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can be transferred into other E. coli strains (14,15) permeable to a variety of hydrophobic and high-molecular-weight substances (up to 5000 daltons). The complementation system can also be set up in naturally more permeable cells, such as Gram-positive eubacteria , mycoplasma, or S. cerevisiae (16). 6. Selected compounds can finally be checked in vitro, in well-defined biochemical assays of purified DnaK (ATPase activity or re-naturation of denatured luciferase, for example). In parallel, the activities of the eucaryotic HSP70 and HSC70 in the presence of these compounds can also be tested. As potential antibacterial molecules, these inhibitors should be more active on bacterial DnaK than on the eukaryotic chaperones. However, inhibitors of eukaryotic chaperones could also be very interesting, because of their participation in many pathways in immunology (presentation of the antigen) and in molecular cancer biology (anti-apoptosis factors).

4. Notes 1. The sidB1 mutation is an extragenic suppressor present in the strain BB1553, in addition to the disrupted dnaK gene. It prevents the excessive synthesis of heatshock proteins that occurs in the absence of DnaK, the negative regulator of the heat-shock response in prokaryotes. It is a mutant allele of rpoH, which encodes the heat-shock-specific transcription factor  32 . The point mutation (Thr252Met) in sidB1 affects the activity rather than the cellular concentration of  32 and suppresses the major cellular defects of the dnaK52 mutant by reducing the elevated level of expression of heat-shock proteins (9). 2. Strains AL19 and BB1553 (dnaK) are thermosensitive for growth on LB plates at 44 °C and 38 °C, respectively, whereas dnaK+ strains AL26 and MC4100 grow on LB plates at 44 °C. BB1553 (cmR ) grows on LB plates containing 25 μ g/mL chloramphenicol at 30 °C. 3. Strain JM83 (lacZM15, dnaK52::cmR ) is chloramphenicol-resistant, temperature-sensitive (at 40 °C), and cold-sensitive (at 20 °C), as expected. When transformed to kanamycin resistance with plasmid pWSK129, it is unable to sustain -complementation after streaking on LB plates containing IPTG, X-Gal, and kanamycin, since all the isolated colonies are white after 48 h at 30 °C. However, the high-cell-density portion of the streak is pale blue (see Fig. 5A of (8)), and therefore it is important to examine the lac phenotype at the level of isolated colonies and not at high cell density. 4. Strain TR20 (dnaK756-ts) grows normally in LB medium at 37 °C and also at 40 °C, although a slight inhibition of growth is observed at 40 °C. -complementation is abolished at these temperatures but is normal at 30 °C, the permissive temperature. 5. In the case of a pulse-chase experiment, labeling is performed for 30 s (with [5-3 H] uridine Amersham (TRK178) isotopically undiluted), followed by a chase with 0.5 mM unlabeled uridine. 6. It is assumed that the mutant DnaK756 protein present in strain AL19 is thermosensitive and inactive only when synthesized at 43 °C, but that the portion synthesized

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at 30 °C is still active and thermoresistant at 43 °C. Therefore, a certain period of time (1.5 h) is needed for its dilution and/or decay in bacteria shifted to the non-permissive temperature. 7. Strains containing pUC18 are grown in LB medium containing 100 μg/mL ampicillin, and those containing pWSK129 in the same medium containing 25 μg/mL kanamycin. 8. In the case of strains harboring pUC18, addition of IPTG to the culture medium is unecessary, because of the titration of the LacI repressor by the numerous pUC18borne lac operators. pUC18 is a high-copy-number plasmid with 100 to 200 copies per cell.

References 1 Alix, J. H. (1993) Extrinsic factors in ribosome assembly. In The Translational 1. Apparatus: Structure, Function, Regulation, Evolution (K. H. Nierhaus et al., eds.), Plenum, New York, pp.173–184. 2 Culver, G. M. (2003) Assembly of the 30S ribosomal subunit. Biopolymers 68, 2. 234–249. 3 Alix, J. H., and Guérin, M. F. (1993) Mutant DnaK chaperones cause ribosome 3. assembly defects in E. coli. Proc. Natl. Acad. Sci. USA 90, 9725–9729. 4 Sbaï, M., and Alix, J. H. (1998) DnaK-dependent ribosome biogenesis in E. coli: 4. Competition for dominance between the alleles dnaK756 and dnaK+ . Mol. Gen. Genet . 260, 199–206. 5 El Hage, A., Sbaï, M., and Alix, J. H. (2001) The chaperonin GroEL and other 5. heat-shock proteins, besides DnaK, participate in ribosome biogenesis in E. coli. Mol. Gen. Genet. 264, 796–808. 6 El Hage, A., and Alix, J. H. (2004) Authentic precursors to ribosomal subunits 6. accumulate in E. coli in the absence of functional DnaK chaperone. Mol. Microbiol. 51, 189–201. 7 Alix, J. H. (2004) The work of chaperones. In Protein Synthesis and Ribosome 7. Structure (K. H. Nierhaus.and D. N. Wilson, eds.), Wiley-VCH Verlag, New York, pp. 529–562. 8 Lopes Ferreira, N., and Alix, J. H. (2002) The DnaK chaperone is necessary for 8. -complementation of -galactosidase in E. coli. J. Bacteriol. 184, 7047–7054. 9 Bukau, B., and Walker, G. C. (1990) Mutations altering heat shock specific subunit 9. of RNA polymerase suppress major cellular defects of E. coli mutants lacking the DnaK chaperone. EMBO J. 9, 4027–4036. 10 Miller, J. H. (1972) Experiments in Molecular Genetics. Cold Spring Harbor 10. Laboratory Press, New York. 11 Neidhardt, F. C., Bloch, P. L., and Smith, D. F. (1974) Culture medium for 11. enterobacteria. J. Bacteriol. 119, 736–747.

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12 Yanisch-Perron, C., Vieira, J., and Messing, J. (1985) Improved M13 phage 12. cloning vectors and host strains: Nucleotide sequences of the M13mp18 and pUC19 vectors. Gene 33, 103–119. 13 Wang, R. F., and Kushner, S. R. (1991) Construction of versatile low-copy-number 13. vectors for cloning, sequencing and gene expression in E. coli. Gene 100, 195–199. 14 Rida, S., Caillet, J., and Alix, J. H. (1996) Amplification of a novel gene, sanA, 14. abolishes a vancomycin-sensitive defect in E. coli. J. Bacteriol. 178, 94–102. 15 Shapiro, S., and Baneyx, F. (2002) Stress-based identification and classification 15. of antibacterial agents: Second-generation E. coli reporter strains and optimization of detection. Antimicrob. Agents Chemother. 46, 2490–2497. 16 Abbas-Terki, T. and Picard, D. (1999) -Complementation of -galactosidase: An 16. in vivo model substrate for the molecular chaperone heat-shock protein 90 in yeast. Eur. J. Biochem. 266, 517–523.

8 Assays for the Identification of Inhibitors Targeting Specific Translational Steps Letizia Brandi, John Dresios, and Claudio O. Gualerzi

Summary While bacterial protein synthesis is the target of about half of the known antibiotics, the great structural-functional complexity of the translational machinery still offers remarkable opportunities for identifying novel and specific inhibitors of unexploited targets. We designed a knowledge-based in vitro translation assay to identify inhibitors selectively targeting the bacterial or the yeast translational apparatus, preferentially blocking the early steps of protein synthesis. Using a natural-like, “universal” model mRNA and cell-free extracts prepared from Eschericha coli, Saccharomyces cerevisiae, and HeLa cells, we were able to translate, with comparable yields in the three systems, the immunogenic peptide encoded by this “universal” mRNA. The immuno-enzymatic quantification of the translated peptide in the presence of a potential inhibitor can identify a selective bacterial or fungal inhibitor inactive in the human system. When applied to the high-throughput screening (HTS) of a library of approximately 25,000 natural products, this assay led to the identification of two novel and specific inhibitors of bacterial translation.

Key Words: translation inhibitors; bacterial translation initiation; eukaryotic translation.

1. Introduction The translational apparatus of both bacteria and lower eukaryotes, such as pathogenic protozoa and fungi, is an ideal target for the identification of new anti-infective drugs. In fact, protein synthesis is a vital, multi-target process involving the largest and most complex ribozyme found in nature (i.e., the ribosome) in addition to over 100 RNA and protein molecules. The mechanism of protein synthesis is essentially the same in all kingdoms of life and relies From: Methods in Molecular Medicine, Vol. 142: New Antibiotic Targets Edited by: W. Scott Champney © Humana Press Inc., Totowa, NJ

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on a large number of molecular components, most of which have a common ancestral origin. Nevertheless, prokaryotic translation and eukaryotic translation are significantly different, at least for some of the components involved and for the mechanistic aspects of some steps of their pathways (1) so as to confer at least kingdom-specificity to a large number of antibiotics targeting the translational apparatus. Furthermore, protein synthesis can be easily reproduced in vitro as the whole process or dissected into its single steps, which can be individually studied. Finally, the large body of information available concerning both structural and functional aspects of the translational apparatus, including the recently elucidated high-resolution 3D structure of the bacterial ribosome alone (2–8) or in complex with antibiotics (9), ensures that identification, characterization, improvement, and rational design of inhibitors can be further pursued as a truly knowledge-based research. The success of the translation apparatus as an antibiotic target is demonstrated by the fact that it represents the target of approximately half of all known antibacterial agents (10–12). In spite of this, the structural-functional complexity of the translational pathway and of its components still offers remarkable opportunities for identifying new bacterial inhibitors specific for unexploited targets such as, for instance, the translation initiation step. On the other hand, the translational apparatus of pathogenic lower eukaryotes (e.g., protozoa and fungi), unlike that of bacteria, has rarely been found as a target for antiinfective drugs active against these organisms. Thus, the simple identification of a translation inhibitor selectively active on lower and not on higher eukaryotic systems would be novel and important for the control of the infectious diseases caused by these organisms. We have designed a model mRNA that displays all the relevant characteristics of a natural bacterial mRNA and is therefore able to drive an initiationand termination-dependent translation in the presence of a bacterial cell extract. The same mRNA has then been appropriately modified to direct eukaryotic translation in the presence of either yeast or mammalian cell-free extracts, thus yielding a “universal” mRNA capable of programming, with comparable efficiency, translational systems derived from three different cellular sources (Eschericha coli, Saccharomyces cerevisiae, and human HeLa cells). Thus, the same peptide synthesized in the three systems can be detected by the same immune-enzymatic reaction, thereby allowing a direct comparison of the inhibitory power of either natural or synthetic compounds in prokaryotic, lower and higher eukaryotic systems. The main characteristics of the “universal” model mRNA are described in Section 1.1.

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When adapted to the high-throughput-screening (HTS) format and applied to the screening of a library consisting of several thousands of natural products, this approach led to the identification of two novel inhibitors of bacterial protein synthesis such as GE81112 (13,14) and GE82832 (15) as well as of other bacteria-specific and yeast-specific inhibitors (unpublished results). 1.1. Characteristics of the “Universal” Model mRNA The model mRNA designated “27IF2Cp(A) mRNA,” schematically illustrated in Fig. 1, contains the relevant characteristics of mRNA naturally occurring in prokaryotes and eukaryotes. Hence, it can be regarded as a universal mRNA that can be used to drive translation in cell-free systems derived from bacterial and low and high eukaryotic cells (i.e., E. coli, S. cerevisiae, and HeLa cells). The main features of the “27IF2Cp(A) mRNA” are the following: 1. Important for prokaryotic translation is the presence of a translation initiation region (T.I.R.) (16) comprising a short Shine and Dalgarno (SD) sequence (AGGU), a 9-nucleotide spacer between the SD and the initiation codon, and a canonical AUG initiation triplet. 2. Important for the eukaryotic translation (17,18) are the sequence context neighboring the AUG start triplet that contains an A at position –3 and a U at position +4, the length of the 5’UTR (which is 74 nucleotides long); the AU-rich nature of the leader region was designed to lack secondary structures; the 3‘UTR containing a poly (A) tail of 26 adenosines that allows CAP-independent translation initiation (see Note 1). 3. The coding sequence is composed of the synthetic 027 gene, encoding the 027 peptide, followed by a cleavable linker of 16 amino acids and the natural coding sequence for the 220 amino acids long C-terminal domain of B. stearothermophilus initiation factor IF2. The 027 gene consists of the 002 mRNA sequence (19) encoding for only 3 amino acids (Phe, Ile, Thr) repeated 3.5 times for a total of 35 residues. In contrast to the amino acid composition of the 027 peptide, the linker and the IF2C sequence contain all the natural amino acids with the exception of Trp.

2. Materials 2.1. General Materials 1. Mortar (20 cm diameter). 2. Polystyrene roller bottles (Corning 850 cm2 ). 3. Dialysis tubing (Spectrum Laboratories, Inc.) with 3,500 Da molecular weight cut-off.

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L. Brandi et al. 5’- gggaauucaaaaauuuaaaaguuaacagguauacauacu AUG UUU ACG AUU M F T I ACU ACG AUC UUC UUU ACG AUC UUC UUU ACG AUU ACU ACG AUC UUC T T I F F T I F F T I T T I F UUU ACG AUU ACU ACG AUC UUC UUU ACG AUU ACU ACG AUC UUC UUU F T I T T I F F T I T T I F F ACG GGG CUG GUU CCG CGU GGA UCC UCU AGA GUC GAC CUG CAG GAG T G L V P R G S S R V D L Q E CAG CGG AGC GUG AAA ACG CGC GUC AGC CUC GAC GAU UUG UUC GAA Q R S V K T R V S L D D L F E CAA AUC AAG CAA GGC GAA AUG AAA GAG CUG AAC UUG AUC GUC AAA Q I K Q G E M K E L N L I V K GCC GAU GUU CAA GGA UCG GUU GAG GCG CUG GUU GCC GCC UUG CAA A D V Q G S V E A L V A A L Q AAA AUC GAU GUC GAA GGC GUG CGG GUG AAA AUC AUC CAU GCG GCG K I D V E G V R V K I I H A A GUC GGC GCC AUC ACC GAA UCG GAU AUU UCA CUG GCG ACG GCA UCG V G A I T E S D I S L A T A S AAC GCU AUC GUC AUU GGC UUU AAC GUC CGC CCG GAU GCG AAU GCG N A I V I G F N V R P D A N A AAG CGC GCC GCU GAA UCG GAA AAA GUC GAC AUC CGC CUC CAU CGC K R A A E S E K V D I R L H R AUC AUU UAC AAC GUC AUU GAG GAA AUU GAA GCG GCG AUG AAA GGG I I Y N V I E E I E A A M K G AUG CUC GAC CCG GAA UAC GAA GAG AAA GUC AUC GGC CAA GCG GAA M L D P E Y E E K V I G Q A E GUG CGG CAA ACG UUC AAA GUG UCC AAA GUU GGC ACG AUC GCC GGC V R Q T F K V S K V G T I A G UGC UAU GUC ACC GAC GGC AAA AUC ACC CGC GAC AGC AAA GUG CGC C Y V T D G K I T R D S K V R CUC AUC CGC CAA GGC AUC GUC GUG UAC GAA GGC GAA AUC GAU UCG L I R Q G I V V Y E G E I D S CUC AAA CGG UAU AAA GAG GAU GUG CGC CAA GUG GCG CAA GGA UAU L K R Y K E D V R Q V A Q G Y GAG UGC GGC UUG ACG AUC AAA AAC UUC AAU GAC AUU AAA GAA GGG E C G L T I K N F N D I K E G GAC GUU AUU GAA GCG UAC GUC AUG CAG GAA GUG GCU CGG GCA UGA D V I E A Y V M Q E V A R A ucggcuuugccgcatgcaagcu(A)26 gcuu - 3’

Fig. 1. Sequence of the “universal” 27IF2Cp(A) mRNA and of its product. The Shine–Dalgarno sequence, the initiation, and termination triplets are written in bold and underlined. The 5‘UTR and 3‘UTR are in lowercase letters, and the open reading frames are in uppercase letters with the nucleotides encoding the thrombin-sensitive linker underlined. The amino acid sequence of the 027 peptide is indicated in bold letters. The amino acids of the thrombin-sensitive spacer (uppercase single-letter amino acids) correspond to the underlined nucleotide sequences and are followed by the Bacillus stearothermophilus IF2 C-domain (uppercase amino acids).

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3MM paper 25 mm round filters (Schleicher & Schuell). Oligo d(T) cellulose Type 7 (Amersham). Sephadex G-25 Fine (Amersham). Unwashed glass beads (425–600 μm, Sigma) (see Note 2). Slot-blot apparatus (BioRad) with nitrocellulose membrane (0.45 μm) (Schleicher & Schuell). Protease Inhibitor Cocktail for Fungal and Yeast extracts (Sigma) and complete EDTA-free Proteinase Inhibitors Cocktail (Roche). [14 C] Phenylalanine, 93.8 mCi/mmol (Amersham). Ultima Gold scintillation liquid (Packard). 9F11 monoclonal antibody (Areta International). Anti-mouse Horseradish-Peroxidase-conjugated (Amersham).

2.2. Stock Solutions 1. Ribonucleotide triphosphates, Na salts (Roche) are dissolved in DEPC-treated H2 O (see Note 3) at 0.1 M. The pH is adjusted to 7.1–7.5 (at room temperature) by addition of 5 N KOH. The solutions are divided in aliquots and stored at –20 °C. 2. Spermidine trihydrochloride and DTT are dissolved in DEPC-treated H2 O to 1 M, divided in aliquots and stored at –20 °C. 3. Na acetate is dissolved in sterile water to 3 M, adjusted to pH 5.2 with glacial acetic acid, sterilized by autoclaving, divided in aliquots, and stored at –20 °C. 4. 150 mL of 50% glucose is made by adding 100 mL of water to 75 g of powder. The powder will go into solution upon autoclaving. 5. E. coli MRE600 tRNA is dissolved in sterile water to 100 mg/mL and bovine liver tRNA to 6 mg/mL. The solutions are divided in aliquots and stored at –20 °C. 6. Benzamidine is dissolved in sterile water to 0.1 M and stored at 4 °C up to a few months. 7. Phenylmethylsulfonyl fluoride (PMSF) (Caution: very toxic upon inhalation and contact with skin) is dissolved in ethanol to 0.1 M and stored up to a few months at 4 °C. 8. Phospho enol pyruvate (0.4 M): The powder is resuspended in 1 N KOH. Sterile water is added, and the pH is adjusted to 7.0 with 1 N (or more concentrated) KOH and brought to the final volume with sterile water. 9. Amino acids mixture: starting from the stock solution (in sterile water) of each amino acid (0.1 M all amino acids but for glu, asn, trp, asp, and tyr, which are 0.05 M). A solution of the desired composition is made by mixing the appropriate amino acids and diluting to a final concentration of 2.5 mM each in 100 mM Tris-HCl, pH 7.7. The final pH should be around 7. 10. 10-Formyl-tetrahydrofolate (10-formyl THF): 10 mg of 5-formyl-tetrahydrofolate (5- formyl THF) are dissolved in 1 mL of 0.1 M HCl that has been flushed extensively with N2 . Store the well-closed vial at 4 °C overnight to obtain a yellow suspension of 5,10-methenyl THF. This suspension is agitated and an aliquot

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L. Brandi et al. withdrawn for spectrophotometric determination of the concentration (6 mM solution = 85 OD355nm ). The expected 5,10-methenyl THF concentration is approximately 20 mM. Since the 10-formyl THF is stable for only a few weeks, it is convenient to store the cyclic methenyl THF, which is stable for several weeks at –20 °C, and prepare fresh 10-formyl THF as needed by hydrolyzing the cyclic methenyl THF. For this, dilute the stock solution in 100 mM Tris-HCl, pH 8.0, extensively flushed with N2 and supplemented with 100 mM of 2-mercaptoethanol. After 15 min incubation at 20 °C, divide the solution in aliquots and store at –20 °C. 3-Aminoethylcarbazole (AEC) stock solution: Dissolve 0.4 g of 3-aminoethyl carbazole (Caution: toxic) in 100 mL of N,N dimethyl formamide. Staining solution for the Western blot: 0.67 mL of AEC stock solution are diluted in 10 mL 0.1 M Na acetate, pH 5.2. To start the peroxidase reaction, add 10 μL of 30% H2 O2 . Creatine phosphate (CP): Dissolve to 1 M in sterile water, and store in aliquots at –20 °C. Creatine phospho kinase (CPK): Dissolve in 30 mM Hepes-KOH, pH 7.4, and 50% glycerol to 5 mg/mL final concentration, and store in aliquots at –20 °C. Ethidium bromide (Caution: possible carcinogen) stock solution: 10 mg/mL in H2 O. Store at room temperature in a dark bottle. Before use, dilute to 0.5 μg/mL with H2 O. TCA stock solutions: Dissolve 100 g of TCA in 100 mL of water. Dilute to 10% and store at 4 °C; dilute to 5% and store at room temperature.

2.3. Buffers 1. Binding buffer: 20mM Tris-HCl, pH 7.5, 500 mM NaCl, 1mM EDTA, pH 8.0. Sterilized by autoclaving. 2. Wash buffer: 20 mM Tris-HCl, pH 7.5, 100 mM NaCl, 1mM EDTA, pH 8.0. Sterilized by autoclaving. 3. Elution buffer: 20 mM Tris-HCl, pH 7.5, 1mM EDTA, pH 8.0. Sterilized by autoclaving. 4. Buffer A: 10mM Tris-HCl, pH 7.7, 10mM Mg acetate, 60mM NH4 Cl, 10% glycerol. Sterilized by autoclaving. Just before use, make 0.2 mM benzamidine, 0.2 mM PMSF, and 1 mM DTT. 5. Bacterial grinding/extraction buffer: Buffer A containing 0.5 g/L bentonite. Sterilized by autoclaving. 6. Breaking buffer: 30 mM Hepes-KOH, pH 7.4, 100 mM K acetate, 2 mM Mg acetate, 20% glycerol. Sterilized by autoclaving. Add mannitol, 85 g/L, and just before use, make 2 mM DTT and 0.5 mM PMSF.

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7. MC buffer: 10 mM Hepes-KOH, pH 7.6, 10 mM K acetate, 0.5 mM Mg acetate, sterilized by autoclaving and supplemented before use with 5 mM DTT and 1X Complete EDTA-free Proteinase Inhibitors Cocktail (see Note 4). 8. Phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 10mM Na2 HPO4 , 2 mM KH2 PO4 . Adjust the pH to 7.4 with HCl and sterilize by autoclaving. 9. 5X Tris Borate EDTA (TBE): 450 mM Tris, 450 mM borate, 10mM EDTA, pH 8 (see Note 5). 10. RNA loading buffer: 98% (v/v) formamide, 10 mM EDTA, pH 8, 1 mg/mL bromophenol blue, and 1 mg/mL xylene cyanol FF.

2.4. Cells and Growth Media 1. E. coli MRE600 was grown in LB medium: tryptone, 10 g/L, yeast extract, 5 g/L, NaCl, 5 g/L, and separately autoclaved glucose (final concentration: 0.5%). 2. S. cerevisiae SKQ 2M was grown in YPD medium: yeast extract, 10 g/L, peptone, 20 g/L, glucose, 20 g/L. 3. Human HeLa S3 (ATCC CCL-2.2), a clonal derivative of the parent HeLa (ATCC CCL-2) line adapted to grow in suspension and therefore more suitable for production of large biomass, was maintained in Minimum Essential Medium Eagle Joklik Modification, supplemented with 10% Fetal Bovine Serum as suspension cultures in Polystyrene Roller Bottle.

2.5. Preparation of the Model mRNA 2.5.1. For in vitro Transcription To prepare the model mRNA, the following materials are required: 1. DEPC-treated water. 2. Ribonucleotide triphosphates, ATP, GTP, UTP, CTP, Na salts (Roche). 3. BSA RNase-free, 20 mg/mL (Roche), and RNasin Ribonuclease Inhibitor, 40 U/mL (Promega). 4. 1 M Tris-HCl, pH 8.0, 1 M MgCl2 , 1 M spermidine trihydrochloride, and 1 M DTT. 5. T7 RNA polymerase (BioLabs).

2.5.2. For mRNA Purification 1. For purification by oligo d(T) cellulose chromatography: binding buffer, wash buffer, elution buffer in addition to ethanol and 3M Na acetate, pH 5.2 for precipitation. 2. For purification by LiCl precipitation: 5 M LiCl, ethanol, and 3M Na acetate, pH 5.2.

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2.5.3. Quality Control of mRNA For the quality control of the transcript, evaluated by PAGE-UREA electrophoresis, the following materials are required: 1. 2. 3. 4.

5X TBE. 6% acrylamide/Bis-acrylamide/7M urea gel. RNA loading buffer. Ethidium bromide staining solution.

2.6. Preparation of E. coli Cell Extract 1. 2. 3. 4. 5. 6.

Alumina. Mortar and pestle. DNase (RNase-free) (Roche). Bacterial grinding/extraction buffer. Buffer A. Dialysis tubing.

2.7. Preparation of S. cerevisiae Cell Extract 1. 2. 3. 4.

Breaking buffer (with and without mannitol). Glass beads. Protease Inhibitor Cocktail for fungal and yeast extracts. Sephadex G-25 (Fine).

2.8. Preparation of HeLa Cell Extract 1. Tight-fitting Dounce homogenizer. 2. MC buffer. 3. 1X Complete EDTA-free Proteinase Inhibitors Cocktail.

2.9. Bacterial in vitro Translation (Radioactive Product) 1. All the components for in vitro translation are dissolved or diluted using sterile materials (disposable sterile plastic ware or glassware sterilized in oven at 180 °C for 8 h) and DEPC-treated water or water-autoclaved 3–4 times. For buffers and salt solutions, it is preferred to resterilize by autoclaving the final solutions; temperature-sensitive solutions are sterilized by filtration through 0.2 μm nitrocellulose membranes. 2. Bacterial cell extract. 3. Universal model mRNA. 4. 1 M Tris-HCl, pH 7.7. 5. 1 M Mg acetate. 6. 4M NH4 Cl. 7. 1M DTT.

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100 mg/mL tRNA (E. coli). 100 mM ATP. 100 mM GTP. 0.4 M PEP. 2.5 mM amino acids mixture minus Phe. 6 mM 10-formyl THF. [14 C] phenylalanine. 10 mM phenylalanine. 3MM filter paper discs. 10% TCA. 5% TCA. Ethanol-ether 1:1. Ether. Ultima Gold scintillation fluid.

2.10. Bacterial in vitro Translation (Non-radioactive Product) 1. This reaction requires the same components listed in Section 2.9 with the exception that non-radioactive phenylalanine is used in place of [14 C] phenylalanine. 2. The translation product is filtered on 0.45 μm nitrocellulose membranes using a slot-blot apparatus. 3. BSA. 4. 9F11 monoclonal antibody. 5. Tween 20. 6. Anti-mouse horseradish-peroxidase-conjugated antibody. 7. Staining solution for Western blot.

2.11. Yeast in vitro Translation (Radioactive Product) The components for the in vitro yeast translation are prepared using the same precautions used for the bacterial translation. 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

Yeast cell extract. 1 M Hepes-KOH, pH 7.4. 2.5 M K acetate. 1 M Mg acetate. 1 M DTT. 100 mM ATP. 100 mM GTP. 1 M CP. 5 mg/mL CPK. 40 U/mL RNase inhibitor. 2.5 mM amino acid mixture minus Phe.

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12. [14 C] phenylalanine. 13. 10 mM phenylalanine.

The quantification of the product requires the same materials listed for bacterial translation. 2.12. Yeast in vitro Translation (Non-radioactive Product) This reaction requires the same components and materials listed in Section 2.1 with the exception that non-radioactive phenylalanine is used in place of [14 C] phenylalanine and product detection requires the same materials listed in Section 2.10. 2.13. Human in vitro Translation The components and materials required for in vitro translation with HeLa cell extracts and for immunological product detection are the same as those listed in Section 2.12 with the exception that human cell extract and Hepes-KOH, pH 7.6, are used. In addition, this test requires 1 M spermidine and 6 mg/mL bovine liver tRNA. 3. Methods 3.1. In vitro Transcription of Model mRNA The quality of the mRNA is a key element in determining the quality of the results obtained in the translation tests. Thus, special care should be taken in the preparation and purification of the mRNA. The main characteristics of the universal mRNA 27IF2Cp(A) are described in Section 1.1.The DNA template used for the in vitro transcription is present as an insert in a plasmid (pTZ18UmRNA). For the transcription reaction it is possible to use either a DNA template prepared by PCR amplification of the relevant coding sequence or the HindIII-linearized plasmid. The mRNA produced by transcription can be purified (see Note 6) either by column chromatography on oligo (dT) cellulose (Section 3.1.2.1) or by LiCl precipitation (Section 3.1.2.2). Disposable sterile plastic ware as well as glassware oven-sterilized for 4 h at 180 °C after rinsing with DEPC-treated water is used in the preparative transcription of the mRNA. 3.1.1. Transcription Reaction 1. The reaction mix (typically 12 mL) consists of 0.02 μM DNA template, 40 mM Tris-HCl, pH 8.1, 20 mM MgCl2 , 10 mM spermidine, 5 mM DTT, 0.1 mg/mL BSA, 0.05 U/μL RNase inhibitor, 0.004 U/μL PPase, 3.75 mM each ribonucleotide triphosphates, and 1.2–1.7 U/μL T7 RNA polymerase. 2. The reaction mix is incubated for 2–3 h at 37 °C (see Note 7).

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3.1.2. mRNA Purification 3.1.2.1. Purification by oligo-dT Cellulose Chromatography 1. Resuspend (typically, 5 g) oligo-(dT) cellulose in DEPC-treated water and autoclave at 120 °C for 20 min. 2. Pack resin into a small glass column (2.2 cm × 6 cm) and equilibrate with 10 column volumes of Binding buffer. 3. At the end of the transcription reaction, adjust the NaCl concentration of the mix to 0.5 M by addition of 5 M NaCl and load the sample onto the column. Collect the flowthrough and repeat the loading. 4. Wash the column with 10 column volumes of Binding buffer followed by 5 volumes of Wash buffer. 5. Elute the mRNA with 3 column volumes of Elution buffer. Collect fractions corresponding to approximately 1/7th the column volume (see Note 8). 6. Dilute each fraction 1:20 and determine A260 . 7. Pool the fractions corresponding to peak of highest A260 . Add 0.1 volume of 3 M Na acetate (pH 5.2) and 2.5 volumes of cold absolute ethanol. Store at –20 °C for at least 30 min. 8. Centrifuge 30 min at 12K rpm 12,000Xg at 4 °C (Sorvall, SA600 rotor). 9. Wash pellet with 70% ethanol. Air-dry pellet. 10. Resuspend RNA in small volume of DEPC-treated water. Determine concentration spectrophotometrically (1A260 = 40 μg/mL) (see Note 9).

3.1.2.2. Purification by LiCl Precipitation 1. At the end of the transcription reaction, add to the reaction mix an equal volume of sterile 5 M LiCl. Keep on ice for 30 min. 2. Centrifuge 10 min at 12K rpm (12,000Xg) (Sorvall SA600 rotor) at room temperature (see Note 10). 3. Rinse the pellet with 70% ethanol, resuspend it in DEPC-treated water, and add 0.1 volume of 3 M Na acetate (pH 5.2) and 2.5 volumes of cold absolute ethanol. Store at –20 °C for at least 30 min. 4. Centrifuge 10 min at 12K rpm (12,000Xg) (Sorvall SA600 rotor) at 4 °C. Repeat Steps 3 and 4 at least twice. 5. Air-dry the last pellet. Resuspend RNA in a small volume of DEPC-treated water. Determine concentration spectrophotometrically (1 A260 = 40 μg/mL).

3.1.2.3. Analysis of the mRNA Quality by PAGE-Urea 1. To check the success of the transcription reaction, remove 5 μL of reaction mix at the beginning and end of the transcription reaction and add to 5 μL of sample buffer.

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2. Incubate at 65 °C for 5 min and load on a 6% acrylamide-7M urea gel. To check the quality of the purified mRNA, 100–200 μg of RNA are analyzed on the same gel. Run the electrophoresis with 1X TBE running buffer.

3.2. Preparation of E. coli Cells and Cell-Free Extract The E. coli cell-free extract (S30) fraction is prepared from cells grown in a fermenter to early-mid log phase, harvested, and stored at –80 °C. The frozen cells are disrupted by grinding with alumina. The S30 fraction, obtained by centrifugation of the crude cell extract to remove membranes and other debris, is dialyzed and stored at –80 °C. Normally, one obtains 1 mL cell extract per gram of cell paste. The activity of the S30 extract is evaluated in a dose-response experiment. 3.2.1. Growth of E. coli Cells 1. Inoculate 15 L of LB medium containing separately sterilized glucose (f.c. 0.5%) with 300 mL of an overnight culture (A600  4) of E. coli MRE600 cells. Incubate at 37 °C with vigorous aeration to A600  1.2. 2. Harvest the cells by continuous-flow centrifugation at 42K rpm (CEPA laboratory LE centrifuge). 3. Resuspend the cells, and wash three or four times with buffer A. 4. Rapidly freeze and store final cell pellet at –80 °C. The average yield is 35–45 g of washed cells per fermentation.

3.2.2. Preparation of E. coli Cell-Free Extract 1. Grind 50 g frozen E. coli cell paste for approximately 25 min with 75 g pre-cooled alumina in a pre-chilled mortar kept on ice. 2. Add DNAse (RNase-free) (2.5 μg/g cells) to the thick cell slurry. 3. Resuspend the cell slurry in 50 mL Bacterial grinding/extraction buffer and gently stir in a beaker for 10 min at 4 °C. 4. Centrifuge the crude cell extract at 12K rpm (12,000X g) for 15 min at 4 °C in an SA600 Sorvall rotor. 5. Discard pellet containing alumina and cell debris and recentrifuge the supernatant at 12K rpm (12,000Xg) for 60 min at 4 °C (SA600 rotor) to obtain the S30 fraction. 6. Dialyze the S30 extract at 4 °C against 40 volumes of buffer A, changing the buffer every 2–3 h for a total of 3 changes. 7. After dialysis, clarify the extract by centrifugation at 12K rpm for 20 min at 4 °C (SA600 rotor). 8. Store S30 in small aliquots (50 μL, 100 μL, 200 μL) at –80 °C (see Note 11).

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3.3. Preparation of Yeast Cells and Cell-Free Extract The growth conditions for S. cerevisiae are similar to those described for E. coli, but the method for rupturing the cells (which involves shaking with glass beads) and preparing the cell extract are quite different. All operations are performed in a cold room and, whenever possible, with incubations in an ice/NaCl mix to maintain a working temperature between –4° and –5 °C. 3.3.1. Growth of S. cerevisiae SKQ 2 M 1. Inoculate 15 L of YPD medium with 200 mL of a saturated culture of S. cerevisiae SKQ 2 M, and incubate at 28–30 °C for approximately 6 h to A660  1. 2. Harvest the cells by continuous-flow centrifugation (CEPA laboratory LE centrifuge). 3. Resuspend the cells, and wash three to four times with Breaking buffer containing mannitol. 4. Freeze and store the final cell pellet at –80 °C.

3.3.2. Preparation of S. cerevisiae Cell Extract 1. Resuspend cells with mild shaking in Breaking buffer with mannitol (2.5 mL/g cells) supplemented with protease inhibitor cocktail. 2. Distribute the resuspended cells in 50 mL Sorvall centrifuge tubes (3–4 g cells per tube). Add 3–4 g ice-cold glass beads per mL buffer. Cap the tubes. 3. Shake the tubes by hand, making 120 vertical 50 cm long movements/min followed by a 1 min incubation in ice/NaCl (–5 °C). 4. Repeat Step 3 for a total of 4 times. 5. Centrifuge the extract at 14K rpm 15,000Xg for 20 min at -5 °C (Sorvall, SA600 rotor). Collect with a Pasteur pipette the S30 in the phase formed between the upper lipid layer and the pelleted glass beads (see Note 12). 6. Equilibrate the Sephadex G-25 column (resuspend the resin in water and sterilize it by autoclaving) with 5 volumes of Breaking buffer without mannitol. 7. Load crude S30 extract (6–10 mL) onto a Sephadex G-25 column (1.8 × 26 cm, corresponding to ca. 60 mL) and elute with Breaking buffer without mannitol at a flow rate of 1 mL/min, collecting fractions of 1 mL. 8. Dilute 1:100 a small aliquot of each fraction and determine A260 . Pool the excluded fractions with highest A260 and determine the A260 of the pool. Store in suitable aliquots at –80 °C. (See Note 13.)

3.4. Preparation of HeLa Cells and HeLa Cell Extract In contrast to the bacterial and fungal cell extracts, which are prepared from frozen cells, the preparation of HeLa cell extract must start from freshly harvested cells.

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3.4.1. Growth of HeLa Cells 1. A frozen 1 mL aliquot containing approximately 106 HeLa cells is resuspended in 25 mL fresh Minimum Essential Medium Eagle Joklik Modification supplemented with fetal bovine serum (10%) and, in the presence of 5% CO2 , allowed to grow to a cell density of approximately 3 × 105 cells/mL. 2. As the cells multiply in a suspension culture at 37 °C in Corning Polystyrene Roller Bottles, fresh medium is periodically added to maintain a constant density of 3 × 105 cells/mL. 3. HeLa cells are harvested from 2–3 L culture having reached a density of 3–6 × 105 by centrifugation in GS3 rotor for 3 min at 2000 rpm (2000X g) at 4 °C using 250 mL sterile Falcon tubes.

3.4.2. Preparation of HeLa Cell Extract 1. Resuspend freshly harvested HeLa cell pellet in 20 mL sterile PBS and divide the suspension in two 15 mL Falcon tubes. Centrifuge for 2 min at 2000 rpm (2000X g) at 4 °C (Sorvall SA600). 2. Step 1 is repeated 3 times (see Note 14). 3. Resuspend final pellet in volume of MC buffer equal to volume of cell pellet. Leave on ice 5 min. 4. Transfer resuspended cells to ice-cold, tight-fitting Dounce homogenizer. Break cells with 15–18 strokes, pausing a few seconds between each stroke to avoid heating. 5. Centrifuge the homogenate at 10,500 rpm 11,000Xg for 5 min at 4 °C (Sorvall SA600 rotor).

3.5. In vitro Translation 3.5.1. In vitro Bacterial Translation Two systems are available to test bacterial translational activity; one is based on the detection of a radioactive product, the other on its immunological detection. The translational systems yielding a radioactive and a non-radioactive product are essentially the same except for the presence of radioactive or non-radioactive phenylalanine, respectively. The first system is faster and very sensitive because the “detection window” to quantify the product can span from a few to several hundred thousand cpm. The non-radioactive translation assay is suitable for systematic HTS assays in which massive use of radioactive materials is problematic. 3.5.2. In vitro Bacterial Radioactive Translation 1. Translation is performed in 25–50 μL reaction mixtures containing 10 mM Tris-HCl, pH 7.7, 7 mM Mg acetate, 100 mM NH4 Cl (see Note 15), 2 mM DTT, 2 mM ATP,

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0.4 mM GTP, 10 mM PEP, 0.025 mg/mL PK, 0.12 mM 10-formyl-tetrahydrofolate, 3 μg /μL tRNA (E. coli), amino acid mixture containing 0.2 mM of all amino acids with the exception of phenylalanine, 9 μM [14 C] phenylalanine, and 36 μM non-radioactive phenylalanine. Optimized aliquots of the S30 cell extract (generally, 2–4 μL/30 μL reaction volume) and of universal mRNA (generally, 1 μM) are added to the reaction mixture (see Notes 16 and 17). The reaction mixture is incubated for 30–60 min at 37 °C, and 20–40 μL aliquots from each tube are spotted on 3 MM paper discs that are immediately dropped into 10% ice-cold TCA. After 30 min, the filters are transferred into 5% TCA and incubated 10 min at 90 °C. After three 5 min washes at room temperature in 5% TCA, the filters are kept in a 1:1 ethyl ether:ethanol mixture for 10 min and finally in ether for an additional 10 min. The filters are then dried and their radioactivity determined by liquid scintillation counting (see Note 18).

3.5.3. In vitro Bacterial Non-radioactive Translation The non-radioactive translation test is performed using the same protocol described in Section 3.5.2 except that 0.045 mM non-radioactive phenylalanine was used. 1. To detect the product of the reaction, 20 μL aliquots are filtered through a nitrocellulose membrane using a slot-blot apparatus. 2. After washing in PBS for 45–60 min, the membrane is blocked for 1.5–2.0 h in PBS containing 3% BSA before being incubated for 3 h in PBS with 0.3% BSA with a suitable dilution of the first antibody (9F11). 3. After three 10 min washes in PBS with 0.05% Tween 20, the membrane is incubated for 1 h in PBS with 0.3% BSA containing a suitable dilution of the second antibody (Anti-mouse HRP-conjugated). 4. After three 10 min washes in PBS with 0.05% Tween 20, the membrane is incubated in 10 mL of staining solution. 5. The reaction of the antibody-conjugated enzyme is stopped in H2 O, and the stained bands are quantified densitometrically.

3.5.4. In vitro Yeast Translation As for the bacterial translation, translational activity in yeast can also be studied measuring the amount of peptide produced using either a radioactive or an immunological test. 3.5.5. In vitro Yeast Radioactive Translation 1. Translation is performed in 25–50 μL reaction mixtures containing 33 mM HepesKOH, pH 7.4, 160 mM K acetate, 3.8 mM Mg acetate (see Note 19), 3.3 mM DTT,

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0.5 mM ATP, 0.1 mM GTP, 30 mM CP, 20 μg/mL CPK, 200 U/mL RNase inhibitor, amino acid mixture containing 200 μM each of all amino acids (minus phenylalanine), 9 μM [14 C] phenylalanine, and 36 μM non-radioactive phenylalanine. 2. Optimized aliquots of the S30 yeast extract (generally 0.5–1 A260 /tube is the best concentration) and of universal mRNA (generally, 0.056 μM) are added to the reaction mixture (following the same procedure described for bacterial translation reported in Note 17). 3. The reaction mixture is incubated for 90–120 min at 25 °C, then 20 μL aliquots from each sample are spotted on 3 MM filters and processed as described for the bacterial system (see Section 3.5.2).

3.5.6. In vitro Yeast Non-radioactive Translation The reaction mixture is prepared and incubated as described in Section 3.5.5 except for the omission of radioactive phenylalanine and the inclusion of 45 μM non-radioactive phenylalanine. The immunodetection and quantification of the translated peptide are performed as described for the bacterial system (see Section 3.5.3). 3.5.7. In vitro Human Translation 1. Translation is performed in 15–25 μL reaction mixtures containing 16 mM HepesKOH, pH 7.6, 75 mM Ka acetate, 2.5 mM Mg acetate, 2mM DTT, 0.8 mM ATP, 0.1 mM GTP, 20 mM CP, 0.1μg/μL CPK, 0.1 μg/μL bovine liver tRNA, amino acid mixture containing 200 μM each of all amino acids including phenylalanine, and 0.1 mM spermidine. 2. Optimized aliquots of the HeLa cell extract (generally, 6–9 μL/15 μL is the best concentration) and of universal mRNA (generally, 0.3 μM) are added to the reaction mixture (following the same procedure described for bacterial translation reported in Note 17). 3. The translation reaction is incubated for 60 min at 30 °C, and 10–25 μL are used for the immunological quantification of the peptide produced, which is carried out as described for the bacterial system (see Section 3.5.3).

4. Notes 1. Using the commercially available chemical capping system, our mRNA can be capped and used to drive a CAP-dependent translation. 2. The unwashed glass beads were washed in 33% HCl (overnight), then washed with plenty of water, and sterilized in an oven at 180 °C for 8 h. 3. DEPC-treated water is obtained by diluting 1:1000 the DEPC (Caution: toxic). After stirring overnight, the solution is autoclaved at 120 °C for 20 min 2–3 times in order to be sure that all the DEPC is completely removed. Since RNases can be permanently denatured by multiple cycles of autoclaving, another way to sterilize water (especially large volumes) is to autoclave it 3–4 times.

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4. Dissolve one tablette of Complete EDTA-free Proteinase Inhibitors Cocktail in 2 mL of sterile water. This solution (25X) is stable at –20 °C for at least three months. Dilute to 1X before use. 5. TBE is usually made and stored as 5X stock solution. Dilute the stock buffer just before use, and make the gel solution and the electrophoresis buffer from the same concentrated solution. 6. Purification by oligo-d(T) cellulose chromatography is more expensive than LiCl precipitation but offers the advantage that the presence of the poly(A) tail at the 3’ end ensures that the purified mRNA is full length. Furthermore, this method is less time-consuming because the eluted mRNA is then ethanol-precipitated, resuspended in DEPC-treated water, and ready to use. The mRNA precipitated by LiCl must be ethanol-precipitated at least three times to remove all the Li + ions, which may inhibit translation, especially in yeast. In general, purification by oligo-d(T) cellulose chromatography is suitable for small mRNA preparations and LiCl precipitation for large preparations. 7. Under our experimental conditions, an average of 48 A260 /mL of transcription reaction, which is equivalent to 1.9 mg of pure mRNA, is obtained. 8. For short time conservation, the column can be left equilibrated in Binding buffer at 4 °C; for longer storage, the DEPC-H2 O rinsed resin can be autoclaved, equilibrated in Binding buffer, and stored at 4 °C. For very long storage, the resin can be dried under vacuum after washing with 50% ethanol and stored at –20 °C. 9. According to the producer indications, 1 g of oligo-(dT) cellulose can bind 90 A260 of poly(A). Thus, 5 g of resin can be used to purify the mRNA produced in 10–12 mL of transcription reaction. 10. After incubation on ice, the LiCl-precipitated mRNA is collected by centrifugation at room temperature. Centrifugation at 4 °C makes the pellet difficult to resuspend. It is also not recommended to incubate the LiCl-precipitated-mRNA at temperatures below 0 °C. 11. The extract is stable at –80 °C for a few years. If needed, it can be frozen and thawed twice without loss of activity, but it is not recommended to freeze and thaw it more than twice. The activity of each S30 preparation is tested by a dose-response translation test to determine the optimal volume to be used in a translation assay. 12. Usually, 8–10 mL of S30 can be obtained from 8 g of yeast cells. 13. The frozen yeast cell paste is usually kept for no longer than one month, while the yeast S30 stored at –80 °C is stable for a longer period of time. 14. It is important to avoid prolonged contact between cells and PBS (20). 15. When calculating the final concentrations of these ions (Mg++ = 7 mM and NH4 + = 100 mM) in the in vitro translation reaction mixture, one must keep in mind that the S30 contains Mg++ and NH4 + . 16. The activity of S30 and mRNA can vary from one preparation to the next so that the optimal amount must be preliminarily tested.

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17. Keep all components of the translation reaction on ice. Prepare a premix consisting of water, ions, energy system, amino acids, and tRNA in an ice-cold tube. Distribute the S30 into the single Eppendorf tubes, and then add the premix and finally the mRNA. The mRNA can be frozen and thawed many times, but use sterile materials and make a fresh dilution for each experiment in order to add an easy-to-pipette aliquot to the rest of the reagents to start the reaction. Transfer the Eppendorf tubes from ice to a 37 °C incubator. Do not freeze leftover S30 more than once. 18. To calculate the yield of product in pmoles, one must determine the specific activity (cpm/pmole) of the Phe in the reaction mixture. This is done by directly spotting 2 μL of the reaction mix onto a prewashed 3MM filter, determining the cpm on the filter, and correlating the cpm with the known amount of Phe present in the 2 μL. From this specific activity (cpm/pmol Phe) and knowing the number of Phe present in the product peptide, one can calculate the pmoles of peptides synthesized in vitro. 19. When calculating the final concentrations of these ions (Mg++ = 3.8 mM and K+ = 160 mM) in the in vitro translation reaction mixture, one must keep in mind that the S30 contains Mg++ and K+ .

Acknowledgments The authors would like to thank Prof. Fabrizio Loreni for the kind gift of the HeLa S3 cell line. References 1 Bottger, E. C., Springer, B., Prammananan, T., Kidan, Y., and Sander, P. (2001) 1. Structural basis for selectivity and toxicity of ribosomal antibiotics. EMBO Rep. 2, 318–323. 2 Ban, N., Nissen, P., Hansen, J., Moore, P. B., and Steitz, T. A. (2000) The 2. complete atomic structure of the large ribosomal subunit at 2.4 A resolution. Science 289, 905–920. 3 Harms, J., Schluenzen, F., Zarivach, R., Bashan, A., Gat, S., Agmon, I., Bartels, 3. H., Franceschi, F., and Yonath, A. (2001) High resolution structure of the large ribosomal subunit from a mesophilic eubacterium. Cell 107, 679–688. 4 Nissen, P., Hansen, J., Ban, N., Moore, P. B., and Steitz, T. A. (2000) The 4. structural basis of ribosome activity in peptide bond synthesis. Science 289, 920–930. 5 Schluenzen, F., Tocilj, A., Zarivach, R., Harms, J., Gluehmann, M., Janell, 5. D., Bashan, A., Bartels, H., Agmon, I., Franceschi, F., and Yonath, A. (2000) Structure of functionally activated small ribosomal subunit at 3.3 Angstroms resolution. Cell 102, 615–623.

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6 Schuwirth, B. S., Borovinskaya, M. A., Hau, C. W., Zhang, W., Vila-Sanjurjo, 6. A., Holton, J. M., and Cate, J. H. (2005) Structures of the bacterial ribosome at 3.5 A resolution. Science 310, 827–834. 7 Wimberly, B. T., Brodersen, D. E., Clemons, W. M. Jr., Morgan-Warren, R. J., 7. Carter, A. P., Vonrhein, C., Hartsch, T., and Ramakrishnan, V. (2000) Structure of the 30S ribosomal subunit. Nature 407, 327–339. 8 Yusupov, M. M., Yusupova, G. Z., Baucom, A., Lieberman, K., Earnest, T. 8. N., Cate, J H, and Noller,H F. (2001) Crystal structure of the ribosome at 5.5 A resolution. Science 292, 883–896. 9 Poehlsgaard, J., and Douthwaite, S. (2005) The bacterial ribosome as a target for 9. antibiotics. Nat. Rev. Microbiol. 3, 870–881. 10 Bottger, E. C. (2006) The ribosome as a drug target. Trends Biotechnol. 24, 10. 145–147. 11 Harms, J. M., Bartels, H., Schlunzen, F., and Yonath, A. (2003) Antibiotics acting 11. on the translational machinery. J. Cell Sci. 116, 1391–1393. 12 Tenson, T., and Mankin, A. (2006) Antibiotics and the ribosome. Mol. Microbiol. 12. 59,1664–1677. 13 Brandi, L., Lazzarini, A., Cavaletti, L., Abbondi, M., Corti, E., Ciciliato, I., 13. Gastaldo, L., Marazzi, A., Feroggio, M., Fabbretti, A., Maio, A., Colombo, L., Donadio, S., Marinelli, F., Losi, F., Gualerzi, C. O., and Selva, E. (2006) Novel tetrapeptide inhibitors of bacterial protein synthesis produced by a Streptomyces sp. Biochemistry 45, 3692–3702. 14 Brandi, L., Fabbretti, A., La Teana, A., Abbondi, M., Losi, D., Donadio, S., 14. et al. (2006) Specific, efficient, and selective inhibition of prokaryotic translation initiation by a novel peptide antibiatic. Proc Natl Acad Sci USA 103, 39–44. 15 Brandi, L., Fabbretti, A., Di Stefano, M., Lazzarini, A., Abbondi, M., and 15. Gualerzi, C. O. (2006) Characterization of GE82832, a peptide inhibitor of translocation interacting with bacterial 30S ribosomal subunits. RNA 12, 1262–1270. 16 McCarthy, J. E., and Gualerzi, C. (1990) Translational control of prokaryotic 16. gene expression. Trends Genet. 6, 78–85. 17 Cigan, A. M., and Donahue, T. F. (1987) Sequence and structural features 17. associated with translational initiator regions in yeast—A review. Gene 59, 1–18. 18 Iizuka, N., Najita, L., Franzusoff, A., and Sarnow, P. (1994) Cap-dependent 18. and cap-independent translation by internal initiation of mRNAs in cell extracts prepared from Saccharomyces cerevisiae. Mol. Cell Biol. 14, 7322–7330. 19 Calogero, R. A., Pon, C. L., Canonaco, M. A., and Gualerzi, C. O. (1988) 19. Selection of the mRNA translation initiation region by Escherichia coli ribosomes. Proc. Natl. Acad. Sci. USA 85, 6427–6431. 20 Carroll, R., and Lucas-Lenard, J. (1993) Preparation of a cell-free trans20. lation system with minimal loss of initiation factor eIF-2/eIF-2B activity. Anal. Biochem. 212, 17–23.

9 SPARK: A New Peptidyl Transferase Activity Assay Alexander S. Mankin and Norbert Polacek

Summary The formation of peptide bonds is the central chemical reaction during protein synthesis and is catalyzed by the peptidyl transferase center residing in the large ribosomal subunit. This active site is composed of universally conserved rRNA nucleosides. The peptidyl transferase center is by far the most frequently used target site of natural antibiotics in the cell. Here we describe a novel, simple, and convenient method to assess peptide bond formation which we named SPARK. The basic principle of SPARK is the use of two reaction substrates that closely resemble the natural tRNA substrates (one is biotinylated and the other carries a tritium label) that become covalently connected during transpeptidation. Formation of this peptide bond then allows capture and direct quantification of the radiolabled product, now joined to the biotin group, using the scintillation proximity assay technology. Binding of the tritiated radioligand to streptavidin-coated beads causes the excitation of the bead-embedded scintillant, thus resulting in the detection of radioactivity. Since no product purification step is required, SPARK is amenable to simple automation, which makes it useful in high-throughput screens of natural or synthetic compound libraries in the search for novel antibiotics.

Key Words: ribosome; antibiotics; peptidyl transferase; high-throughput screening.

1. Introduction All the proteins in the cell are synthesized by ribosomes, which use the genetic information brought in the form of mRNA to polymerize amino acids into functional polypeptides. The ribosome is an evolutionary preferred antibiotic target: More than half of the known natural antibiotics inhibit growth of bacteria by interfering with the functions of the translation apparatus. From: Methods in Molecular Medicine, Vol. 142: New Antibiotic Targets Edited by: W. Scott Champney © Humana Press Inc., Totowa, NJ

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Fig. 1. The principle of SPARK. a. The reaction of peptide bond formation (peptidyl transfer reaction) catalyzed by the ribosome in the process of protein synthesis. In the result of the reaction, the peptidyl residue (light gray circles) is transferred from the P-site bound tRNA to the aminoacyl moiety (dark gray circle) of the A-site bound aminoacyl-tRNA. b. SPARK-Pmn. The tritium-labeled formyl methionine (filled circle) is transferred from the P-site bound formyl-[3 H]Met-tRNAfMet to the aminoacyl residue (open circle) of biot CC-Pmn (biotin moiety is shown as a star). The reaction product

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The ribosome consists of two subunits, large and small. Each of the subunits is built of ribosomal RNA (rRNA) and ribosomal proteins, with rRNA being the main structural and functional component of the ribosome. The key chemical reaction catalyzed by the ribosome is the formation of new peptide bonds that link amino acid residues into polypeptides. In the course of the reaction, the growing polypeptide is transferred from the peptidyl-tRNA bound in the P-site of the ribosome to the aminoacyl-tRNA bound in the ribosomal A-site. This results in the elongation of the polypeptide by one amino acid (Fig. 1). In chemical terms, the reaction involves a nucleophilic attack of the -amino group of aminoacyl-tRNA on the carbonyl carbon atom of the ester bond that links the peptidyl moiety to the 3’-hydroxyl of the 3’ terminal adenine residue of peptidyl-tRNA. The reaction takes place in the active site of the catalytic center of the large ribosomal subunit, which is called the peptidyl transferase center (PTC). A great variety of natural antibiotics and their semisynthetic derivatives bind at the PTC and interfere with peptide bond formation by obstructing correct binding of the PTC substrates, peptidyl-tRNA and aminoacyl-tRNA, or by affecting the orientation of critical nucleotides in the peptidyl transferase active site (1). Clinically important antibiotics such as lincosamides, streptogramins A, oxazolidinones, and others act upon the PTC. Therefore, a significant effort is dedicated to search for newer drugs that would inhibit translation by targeting the PTC, which arguably represents the best evolutionarily and clinically validated antibiotic target. The Scintillation Proximity Assay for Ribosome Kinetics (SPARK) is an assay developed in collaboration with researchers at Pharmacia and Upjohn (presently Pfizer) (2). SPARK provides a simple and efficient way to test multiple compounds for their ability to inhibit peptide bond formation in a highthroughput format. The reaction utilizes beads that carry embedded scintillant (SPA beads, GE-Healthcare Bio-Sciences) and are coated with streptavidin. SPA beads can detect the radioactivity of a tritium-labeled substrate bound on the surface of the beads. One of the substrates used in SPARK assay carries ◭ Fig. 1. (Continued) is captured by streptavidin-coated SPA beads, and the proximity of the tritium source to the bead leads to activation of the bead-embedded scintillant. c. SPARK-2T. The N-biotin-conjugated phenylalanine residue is transferred from the P-site bound biotin-Phe-tRNAPhe to [3 H]Phe-tRNAPhe of the A-site bound aminoacyltRNA. The reaction product is captured by streptavidin-coated SPA beads, and the proximity of the tritium source to the bead leads to activation of the bead-embedded scintillant.

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a biotin tag, while the other is 3 H-labeled. The peptidyl transfer reaction (the reaction of peptide bond formation) leads to a covalent linkage between the two substrates and thus generates a biotinylated and tritium-labeled product that can be monitored directly in the reaction mixture without further purification (Fig. 1b–c). Depending on the experimenter’s needs, the reaction can utilize various tRNA analogs. The activity of a complete (70S) ribosome or of an isolated large (50S) ribosomal subunit can be assayed. 2. Materials 2.1. Ribosomes and S100 Supernatant 1. E. coli ribosomes and ribosomal subunits were prepared from the strain MRE600 or CAN20-12E, essentially as described in (3). A 25 g cell pellet is resuspended in 45 ml buffer A, 20 mM Tris-HCl, pH 7.5, 100 mM NH4 Cl, 10 mM MgCl2 , 0.5 mM EDTA, 6 mM -mercaptoethanol, and lysed by passing the cells twice through a French Press at 18,000 psi. The cell lysate is centrifuged for 15 min at 30,000X g in an SS-34 rotor and the resulting supernatant incubated with 20 units RQ1 DNase (Promega) for 5 min on ice. After another 15 min centrifugation at 30,000X g, the cell lysate is layered on top of a 12 mL 1.1 M sucrose cushion prepared in buffer B: 20 mM Tris-HCl, pH 7.5, 500 mM NH4 Cl, 10 mM MgCl2 , 0.5 mM EDTA employing 32.4 mL Beckman Optiseal polyallomer tubes. Subsequent to a 17 h ultracentrifugation at 100,000X g, the crude post-ribosomal supernatant (S100) is removed and the ribosome pellet-washed with 5 mL buffer C: 50 mM Tris-HCl, pH 7.6, 100 mM NH4 Cl, 10 mM MgCl2 , 6 mM -mercaptoethanol, and finally resuspended in 300 μL buffer C. The concentration of the 70S ribosome preparation is determined by spectrophotometry (1A260 = 24 pmol 70S). The ribosome concentration is adjusted to 24 μM, and the solution is aliquotted, snap-frozen, and stored at –80°C. 2. In our experience, SPARK can be used with ribosomes from other Gram-positive and Gram-negative species (Staphylococcus aureus, Thermus aquaticus, and others (2,4)) without any modifications of the SPARK protocol. 3. The E. coli post-ribosomal supernatant (S100) containing aminoacyl-tRNA synthetases is used for tRNA aminoacylation (5). The crude S100 supernatant from Step 1 is dialyzed three times against 100 volumes of buffer D: 20 mM HepesKOH, pH 7.6, 6 mM MgOAc2 30 mM NH4 Cl, 4 mM 2-mercaptoethanol for 30 min each at 4°C using a membrane with a molecular-weight cut-off of 3.5 kD. The endogenous tRNAs are removed by batch treatment with DEAE-cellulose (SigmaAldrich, St. Louis, MO) (this step also eliminates a substantial amount of cellular RNases). 5 g of cellulose are resuspended in buffer E: 20 mM Hepes-KOH, pH 7.6, 10 mM MgOAc2 500 mM KCl prewarmed to 90°C, and incubated at 90°C for 30 min. The supernatant is decanted, and the procedure repeated twice at 90°C and three times with buffer F: 20 mM Hepes-KOH, pH 7.6, 10 mM MgOAc2

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150 mM KCl at room temperature. The supernatant is removed, and the cellulose matrix is left for at least 2 h at 4°C and subsequently combined with ∼50 mL S100 extract and incubated for 2 h at 0°C under occasional swirling. After centrifugation (10,000X g at 4°C for 30 min in an SS-34 rotor), the supernatant is removed, and the resin is resuspended in 15 mL of buffer F and incubated for 2 h on ice. The resin is pelleted by centrifugation (10,000X g, 4°C, 30 min in an SS-34 rotor), and the supernatant (fraction II) is saved. The elution step is repeated two more times using 15 mL buffer F200: 20 mM Hepes-KOH, pH 7.6, 10 mM MgOAc2 200 mM KCl (fraction III), and finally 15 ml buffer E (fraction IV). The resulting fractions II, III, and IV are individually dialyzed four times against 2 L of buffer D for 45 min at 4°C and centrifuged for 30 min at 30,000X g at 4°C. Typically, fractions II and III show the highest activities in tRNA aminoacylation reactions. The active S100 fractions are stored in aliquots at –80°C.

2.2. tRNAs and mRNAs 1. E.coli MRE600 tRNAPhe and tRNAfMet (Sigma-Aldrich, St. Louis, MO) were dissolved in water, their concentration adjusted to 100 μM, and stored in aliquots at –20°C. 2. The 10X charging buffer for tRNA aminoacylation: 200 mM Hepes-KOH, pH 7.6, 80 mM MgCl2 , 1.5 M NH4 Cl, 40 mM -mercaptoethanol, 20 mM spermidine, and 0.5 mM spermine. 3. The formyl donor (leucovorin) required for preparation of formyl-Met-tRNAfMet (fMet-tRNA) was synthesized by combining 5 mg of folinic acid (Sigma-Aldrich, St. Louis, MO), 400 μL 50 mM -mercaptoethanol, and 44 μL 1 M HCl for 3 h at room temperature (25°C). Aliquots of leucovorin were stored at –20°C. 4. The tritiated amino acids L-[2,6-3 H]-phenylalanine and L-[methyl-3 H]-methionine were from GE-Healthcare Bio-Sciences (Piscataway, NJ). 5. Sulfosuccinamidyl 6-(biotinamido) hexanoate (Catalog # 21335ZZ, Pierce Biotechnology, Rockford, IL) was used for biotinylating the -amino group of aminoacyl tRNAs, when necessary. 6. An RNA oligonucleotide AAGGAGAUAUAACAAUGGGU (Dharmacon) containing a unique Met codon (underlined) or poly(U) (Sigma-Aldrich, St. Louis, MO) was used as mRNA analogs.

2.3. Biotinylated Puromycin Derivatives 1. The 5’-biotin-puromycin derivative used in the original papers describing SPARK (2,4) is no longer available from the commercial sources. However, 5’-biotin-CCpuromycin (Biot CC-Pmn) (Dharmacon RNA Technologies, Lafayette, CO) can be used as a substitute. Biot CC-Pmn is dissolved in water and stored in aliquots at a concentration of 12 μM at –20°C.

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2.4. Reaction Buffers for SPARK 1. SPARK-Pmn reaction buffer (2X): 40 mM Hepes-KOH, pH 7.6, 60 mM MgOAc2 , 300 mM NH4 Cl, 8 mM -mercaptoethanol, 4 mM spermidine, and 0.1 mM spermine. 2. SPARK-2T reaction buffer (2X): 40 mM Hepes-KOH, pH 7.6, 12 mM MgOAc2 , 300 mM NH4 Cl, 8 mM -mercaptoethanol, 4 mM spermidine, and 0.1 mM spermine (6). 3. SPARK stop mix: 0.45 mg SPA beads per 120 μL of 10 mM sodium phosphate buffer, pH 7.4, 2.7 mM NaCl, and 125 mM EDTA (see Note 1).

2.5. SPA Beads and Product Detection 1. Streptavidin-coated SPA beads (catalog # RPNQ0006, GE-Healthcare Bio-Sciences, Piscataway, NJ). The binding capacity of the SPA beads is 127 pmol biotin/mg beads (see Note 2). 2. A 96-well-plate top counter (e.g., Top Count NXT, Perkin-Elmer) is used when SPARK is performed in a 96-well-plate format. 3. A standard liquid scintillation counter (e.g., Beckman LS 6000IC) is used when SPARK is performed in Eppendorf tubes (see Note 3).

3. Methods SPARK can be used for studying catalysis of peptide bond formation (see Note 4) or in screenings (low- or high-throughput) of natural or synthetic compound libraries for new inhibitors of protein synthesis. It has been optimized to be used with a full-size peptidyl-tRNA analog as a donor substrate and a minimal A-site substrate analog, Biot CC-Pmn, as acceptor (SPARK-Pmn) or with full-length peptidyl- and aminoacyl tRNAs (SPARK-2T). 3.1. Preparation of the Peptidyl-tRNA Donor Substrate for SPARK-Pmn 1. Formyl-[3 H]Met-tRNAfMet is prepared in a final volume of 500 μL of 1X charging buffer supplemented with ATP to a final concentration of 3 mM. The reaction contains 5,000 pmol tRNAfMet , 10,000 pmol L-[methyl-3 H]-methionine (6.5 Ci/mmol), 88 μL leucovorin solution (see Note 5), and 50 μL of S100 cell lysate from E. coli (see Note 6). 2. The reaction is incubated for 25 min at 37°C. 3. Aminoacylation is stopped by adding 50 μL 3M NaOAc (pH 5.5) followed by the addition of one reaction volume of water-saturated phenol and shaking for 5 min at room temperature. After separation of the phases by centrifugation in a table-top microcentrifuge (12,000 rpm, room temperature), the aqueous phase is transferred to another tube and re-extracted with an equal volume of phenol/chloroform and then with chloroform.

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4. Formyl-[3 H]Met-tRNAfMet is precipitated by the addition of three volumes of ethanol and incubation for 5 min at –70°C. The RNA pellet is recovered by centrifugation (12,000 rpm, 4 C°), resuspended in water, and stored in aliquots at –80°C. Formyl[3 H]Met-tRNAfMet can be additionally purified by HPLC following the conditions described in (7).

3.2. Monitoring Peptide Bond Formation by SPARK-Pmn 1. 24 pmol 70S ribosomes (1A260 = 24 pmol) are incubated in a final volume of 57.5 μL SPARK-Pmn reaction buffer with 76 pmol synthetic mRNA and 48 pmol formyl-[3 H]Met-tRNAfMet for 10 min at 37°C (see Note 7). 2. The peptidyl transfer reaction is initiated by the addition of 2.5 μL of SPARK-Pmn reaction buffer containing 30 pmol of biot CC-Pmn and performed at 37°C for 15 min (see Note 8). 3. The reaction is terminated by the addition of 2 volumes (120 μL) SPARK stop solution (containing the SPA beads). 4. For best scintillation readouts, incubate for 1 h at room temperature to allow complete binding of the reaction products to SPA beads and full bead settling prior to counting radioactivity (see Note 9).

3.3. Preparation of the Peptidyl-tRNA Donor Substrate for SPARK-2T 1. Phe-tRNAPhe is prepared in a final volume of 500 μL of 1X charging buffer supplemented with ATP to a final concentration of 3 mM. The reaction contains 5,000 pmol tRNAPhe , 10,000 pmol L-phenylalanine, and 50 μL of S100 cell lysate from E. coli (see Note 6). 2. The reaction is incubated for 25 min at 37°C. 3. Aminoacylation is stopped by adding 50 μL of 3 M NaOAc (pH 5.5) followed by the extraction with water-saturated phenol, phenol/chloroform, and chloroform and ethanol precipitation as described in Section 3.1. 4. Biotin-Phe-tRNAPhe is prepared by combining 200 μL of 50 mM NaHCO3 (pH 8.2) containing 1700 pmol Phe-tRNAPhe with 100 μL of a 7 mg/mL solution of sulfosuccinamidyl 6-(biotinamido) hexanoate and incubating 1 h on ice. BiotinPhe-tRNAPhe is then purified by HPLC following the procedure described in (7). HPLC-purified biotin-Phe-tRNAPhe is ethanol-precipitated and dissolved in H2 O (see Note 10).

3.4. Preparation of the Aminoacyl-tRNA Acceptor Substrate for SPARK-2T 1. [3 H]Phe-tRNAPhe is prepared in a final volume of 1 mL of 1X charging buffer supplemented with ATP to a final concentration of 3 mM. The reaction contains 10,000 pmol tRNAPhe , 30,000 pmol L-[3 H]-phenylalanine (6.5 Ci/mmol), and 100 μL of E. coli S100 cell lysate (see Note 6).

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2. The reaction is incubated for 15 min at 37°C. 3. The reaction is stopped by adding 100 μL of 3 M NaOAc (pH 5.5) followed by the extraction with water-saturated phenol, phenol/chloroform, and chloroform and ethanol precipitation as described in Section 3.1. To improve the SPARK reaction performance, [3 H]Phe-tRNAPhe can be further purified by HPLC as described in (7), ethanol-precipitated, and resuspended in H2 O (see Note 11).

3.5. Monitoring Peptide Bond Formation by SPARK-2T 1. 7 pmol 70S ribosomes are incubated with 5.7 pmol biotin-Phe-tRNAPhe and 40 μg of poly(U) in 22 μL SPARK-2T reaction buffer for 15 min at 37°C. 2. The peptidyl transfer reaction is initiated by the addition of 1.3 μL H2 O containing 5.2 pmol [3 H]Phe-tRNAPhe  The reaction is incubated at 37° C for 20 min (see Note 12). 3. The reaction is stopped by the addition of 47 μL SPARK stop solution (containing the SPA beads) (see Note 13).

4. Notes 1. The SPARK stop mix should always be prepared freshly. 2. The SPA beads should be stored at 4°C and should not be frozen. 3. After the SPARK reaction performed in 1.5 mL Eppendorf tubes has been stopped, the reaction mixture does not need to be pipetted into a scintillation vial; the Eppendorf tube can be directly placed into the scintillation vial instead. 4. The kinetics of the peptidyl transferase reaction as measured by SPARK is slower than that catalyzed by the ribosome in the living cell. This can be due to the presence of biotin on one of the reaction substrates. However, all the tested wellcharacterized inhibitors of peptide bond formation readily inhibit the reaction monitored by SPARK (2,8). 5. Before adding leucovorin to the reaction, the pH of the leucovorin solution has to be increased by adding 2 μL of 5 M KOH to a 120 μL aliquot in order to allow complete dissolving. 6. It is important to adjust the final pH of the charging reaction to 7.5 by using 1 M KOH before the S100 addition. 7. In SPARK reactions containing antibiotics, the drug is added after binding of the donor (P-site) tRNA substrate prior to addition of the acceptor (A-site) substrate. 8. In SPARK-Pmn, the reaction with biot CC-Pmn as an acceptor substrate, a plateau in product formation is reached after 30 min of incubation. 9. SPA beads register ∼35% of the radioactivity of the reaction products compared to conventional liquid scintillation counting. The radioactivity of unreacted substrates is essentially not registered, and background radioactivity values (obtained by omitting ribosomes or the biotinylated substrate) are approximately 2 orders of magnitude lower than the measured radioactivity of the reaction products produced in the uninhibited reaction.

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10. Biotin-Phe-tRNAPhe elutes in two distinct peaks from the HPLC C4 column. While both fractions were shown to be biotinylated, only the slower-eluting tRNA fraction is active in SPARK-2T. 11. HPLC purification of [3 H]Phe-tRNAPhe can be substituted with purification by BD-cellulose column chromatography as described in (9). 12. Under these conditions, the reaction proceeds to completion and the entire Phe-[3 H]Phe-tRNAPhe is converted into the dipeptidyl-tRNA product biotin-Phe[3 H]Phe-tRNAPhe . 13. SPARK-2T can also be used with isolated large ribosomal subunits instead of 70S ribosomes. In this version of SPARK-2T, 14 pmol 50S subunits are incubated with 11 pmol biotin-Phe-tRNAPhe in 28 μL of SPARK-Pmn reaction buffer for 10 min at 37°C. The reaction tube is then transferred on ice. 2 μL of H2 O containing 12 pmol [3 H]Phe-tRNAPhe are added, and the reaction is initiated by the addition of 15 μL of cold methanol. The reaction is incubated on ice for 4 h and stopped by the addition of 90 μL of SPARK stop mix (containing SPA beads).

Acknowledgments The authors would like to thank Dean Shinabarger and Steven Swaney (Pharmacia and Upjohn, currently Pfizer) for a fruitful collaboration leading to the development of SPARK. Steven Swaney is additionally thanked for advice on preparing the manuscript. This work was supported by NIH Grant GM 59028 to A.S.M. and by the Austrian Science Foundation (FWF) Grant P18709 to N.P. References 1 Polacek, N., and Mankin, A. S. (2005) The ribosomal peptidyl transferase center: 1. Structure, function, evolution, inhibition. Crit. Rev. Biochem. Mol. 40, 285–311. 2 Polacek, N., Swaney, S., Shinabarger, S. D., and Mankin, A. S. (2002) SPARK—A 2. novel method to monitor ribosomal peptidyl transferase activity. Biochemistry 41, 11602–11610. 3 Moazed, D., and Noller, H. F. (1989) Interaction of tRNA with 23S rRNA in the 3. ribosomal A, P, and E sites. Cell 57, 585–597. 4 Polacek, N., Gaynor, M., Yassin, A., and Mankin, A. S. (2001) Ribosomal peptidyl 4. transferase can withstand mutations at the putative catalytic nucleotide. Nature 411, 498–501. 5 Triana-Alonso, F. J., Spahn, C. M., Burkhardt, N., Rohrdanz, B., and Nierhaus, K. H. 5. (2000) Experimental prerequisites for determination of tRNA binding to ribosomes from Escherichia coli. Methods Enzymol. 317, 261–276. 6 Blaha, G., Stelzl, U., Spahn, C. M., Agrawal, R. K., Frank, J., and Nierhaus, K. H. 6. (2000) Preparation of functional ribosomal complexes and effect of buffer conditions

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on tRNA positions observed by cryoelectron microscopy. Methods Enzymol. 317, 292–309. 7 Erlacher, M. D., Lang, K., Shankaran, N., Wotzel, B., Huttenhofer, A., Micura, R., 7. et al. (2005) Chemical engineering of the peptidyl transferase center reveals an important role of the 2’-hydroxyl group of A2451. Nucleic Acids Res. 33, 1618–1627. 8 Polacek, N., Gomez, M. G., Ito, K., Nakamura, Y., and Mankin, A. S. (2003) The 8. critical role of the universally conserved A2602 of 23S ribosomal RNA in the release of the nascent peptide during translation termination. Mol. Cell 11, 103–112. 9 Robertson, J. M., and Wintermeyer, W. (1981) Effect of translocation on topology 9. and conformation of anticodon and D loops of tRNAPhe . J. Mol. Biol. 151, 57–79.

10 High-Throughput Screening of Peptide Deformylase Inhibitors Kiet T. Nguyen and Dehua Pei

Summary The emergence of bacterial pathogens resistant to current antibiotics has caused an urgent demand for new treatments. Peptide deformylase (PDF) has become an exciting target for designing novel antibiotics. To facilitate the screening of PDF inhibitors, three robust, coupled assays have been developed. The first method couples the PDF reaction with that of formate dehydrogenase. Formate dehydrogenase oxidizes formate into CO2 with a concomitant reduction of NAD+ to NADH, which can be monitored spectrophotometrically. The second method involves Aeromonas aminopeptidase (AAP) as the coupling enzyme and an artificial substrate, f-Met-Leu-p-nitroanilide. The sequential action of PDF and AAP releases p-nitroanilide as a highly chromogenic product. In the third method, f-Met-Lys-7-amino-4-methylcoumarin is used as the substrate. Deformylation by PDF gives an excellent substrate for dipeptidyl peptidase I, which releases the dipeptide Met-Lys and fluorogenic 7-amino-4-methylcoumarin. The combination of these assay methods should meet the needs of most laboratories.

Key Words: antibacterial agents; Aeromonas aminopeptidase; continuous assay; end-point assay; dipeptidyl peptidase I; peptide deformylase; deformylase inhibitor; formate dehydrogenase; N-formyl substrates.

1. Introduction Peptide deformylase (PDF) has emerged as an exciting target for designing novel antibacterial drugs to combat the ever-increasing antibiotic-resistant pathogens (1–4). Bacterial protein biosynthesis starts with N-formylmethionine, and, as a result, all newly synthesized polypeptides carry an N-terminal formyl From: Methods in Molecular Medicine, Vol. 142: New Antibiotic Targets Edited by: W. Scott Champney © Humana Press Inc., Totowa, NJ

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group (5). PDF catalyzes the hydrolytic removal of the N-formyl moiety from the vast majority of these polypeptides. Subsequently, many of these polypeptides undergo further processing by methionine aminopeptidase to produce mature proteins (6). The essentiality of PDF has been demonstrated both by genetic studies (7–9) and by growth inhibition by PDF inhibitors (1–4). PDF is present in all eubacteria and has a highly conserved active site, which contains a tetrahedrally coordinated and catalytically essential divalent metal ion (Fe2+ in most bacteria) (10,11). Although PDF is also present in human mitochondrion, its physiological function in the mitochondrion remains controversial, and specific PDF inhibitors have shown little toxicity to human cells (12). These properties have made PDF an attractive antibacterial drug target. Many potent PDF inhibitors have been reported in recent years, and one of the inhibitors has already advanced into phase I clinical trials for the treatment of upper respiratory tract infections. To facilitate mechanistic study of this enzyme and screening of PDF inhibitors, several robust assays have been developed for PDF. One method monitors the release of formate by formate dehydrogenase (FDH) as the coupling enzyme (Fig. 1a) (13,14). Oxidation of formate into CO2 is coupled with the reduction of NAD+ into NADH, which can be monitored at 340 nm. The second assay method employs Nformylmethionyl-leucyl-p-nitroanilide (f-ML-pNA) as the substrate, and the PDF reaction is coupled with Aeromonas aminopeptidase (AAP). Deformylation by PDF releases ML-p-nitroanilide, which is sequentially hydrolyzed by AAP to release methionine, leucine, and the chromophore p-nitroaniline (Fig. 1b) (15). The third method employs N-formylmethionyl-lysyl-7-amino4-methylcoumarin (f-MK-AMC) as the substrate (16). The PDF reaction produces MK-AMC as its product, which is an excellent substrate for dipeptidyl peptidase I (DPPI). DPPI hydrolysis generates 7-amino-4-methylcoumarin as the chomophore/fluorophore (Fig. 1c). Although each assay method has its strengths and weaknesses, a combination of the three methods should meet most of the assay needs. As mentioned above, native PDFs from most bacteria contain a Fe2+ ion in the active site, which makes them extremely unstable under aerobic conditions (17). A common practice has been to replace the Fe2+ ion with a more stable metal ion such as Co2+ , Ni2+ , or Zn2+ . Both Ni(II)- and Co(II)-substituted PDF variants retain essentially full catalytic activity and are highly stable (11,18,19). They are frequently used in mechanistic studies and PDF inhibitor screening. The Zn(II)-substituted enzyme is very stable but has greatly reduced activity (>100-fold).

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NAD+ NADH

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Fig. 1. Reactions involved in PDF coupled assays. (a) With FDH as coupling enzyme; (b) with AAP as coupling enzyme; and (c) with DPPI as coupling enzyme.

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2. Materials 2.1. Bacterial Strains and Expression Vectors 1. Escherichia coli strain DH5 cells [F− , 80dlac(lacZ)M15 (lacZYA-argF), U169, endA1, recA1, hsdS17 (rK − m K + ), deoR, thi-1, supE44, gyrA96, relA1] for subcloning and construction of expression vectors. 2. E. coli strain BL21 (DE3) cells [F− ,OmpThsdS B (rB − m B − ), gal, dcm, met (DE3)]. This is used for protein overexpression (see Note 1). 3. Plasmid pET22b(+) (Novagen, Madison, WI) as cloning and expression vector to add a histidine tag to the C-terminus of PDF. Clone PDF gene by polymerase chain reaction (PCR) and ligate it into NdeI and XhoI sites of pET22b to construct pET22b-PDF.

2.2. Buffers and Materials for Protein Expression and Purification 1. Luria-Bertani (LB) media: 10 g/L Bacto Tryptone (Difco), 5 g/L yeast extract (Difco), 5 g/L NaCl. Prepare the media in double-distilled H2 O, and autoclave it before use. 2. Minimal media: 5 g/L Bacto casamino acids (acid hydrolysate of casein, Difco), 10.8 g/L K2 HPO4 , 5.5 g/L KH2 PO4 , 10 g/L NaCl in Millipore water with a resistance at dissipation of 18.3 M. Autoclave at 121 °C for 20 min. 3. 1000X Trace metal supplement: 124 mg/L MgSO4  7H2 O, 74 μg/L CaCl2  2H2 O, 20 μg/L MnCl2  4H2 O, 31 μg/L H3 BO3 , 1.2 μg/L (NH4 )6 Mo7 O24  4H2 O, 1.6 μg/L CuSO4 in 0.1 M HCl solution. 4. 100X Nutrient mix: 25% D-(+)-glucose, 10% ammonium sulfate, 20 mg/mL thiamine (Sigma), and 10 mg/mL D-(+)-biotin (Sigma). Sterilize the solution by filtration through a 0.45 μM nitrocellulose membrane filter (Nalgen). 5. Isopropyl--D-thiogalactopyranoside (IPTG) (Sigma) should be prepared fresh at 1000X concentration (100 mM stock) and filtered through the 0.45 μM membrane. 6. Lysis buffer (buffer A): 20 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), pH 7.9, 500 mM NaCl, 5 mM imidazole, 0.1 mg/mL phenylmethylsulfonyl fluoride (PMSF), 40 μg/mL trypsin inhibitor, 100 μg/mL lysozyme, 1% Triton X-100, 0.05% protamine sulfate. All reagents are from Sigma. Prepare fresh buffer. 7. Tris(2-carboxyethyl)phosphine (TCEP) (Aldrich, Milwaukee, WI). 8. Talon resin (Clontech, Mountain View, CA) for cobalt affinity column chromatography. 9. Talon column wash buffer (buffer B): 20 mM HEPES, pH 7.9, 500 mM NaCl, 5 mM imidazole. Store at 4 °C. 10. Talon column elution buffer (buffer C): 20 mM HEPES, pH 7.9, 150 mM NaCl, 60 mM imidazole. 11. Dialysis buffer (buffer D): 20 mM HEPES, pH 7.0, 150 mM NaCl. Store at 4 °C. 12. Ni-NTA His-bind Resin (Novagen).

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13. Ni-NTA His-bind buffer (buffer E): 50 mM sodium phosphate, pH 8.0, 300 mM NaCl, 10 mM imidazole. Store at 4 °C. 14. Ni-NTA elution buffer (buffer F): 50 mM sodium phosphate, 300 mM NaCl; 125 mM imidazole. Store at 4 °C. 15. Enzyme assay buffers. 10X FDH assay buffer (buffer G): 500 mM sodium phosphate, pH 7.0; 10X AAP assay buffer (buffer H): 500 mM HEPES, pH 7.0, 1.5 M NaCl; 10X DPPI assay buffer (buffer I): 500 mM HEPES, pH 7.0, 1.5 mM NaCl, 50 mM DTT. Store at room temperature. 16. Anion exchange buffers. Low-salt buffer (buffer J): 20 mM 2-[Nmorpholino]ethanesulfonic acid (MES), pH 6.0, 10 mM NaCl. High-salt buffer: 20 mM MES, pH 6.0, 1.0 M NaCl. Store at 4 °C. 17. Enzyme dilution buffer (buffer K): 50 mM HEPES, pH 7.0, 100 μg/mL bovine serum albumin. 18. MonoQ HR 5/5 FPLC anion exchange column (Pharmacia).

2.3. Protein Quantitation and Storage 1. Protein assay dye reagent concentrate (Bio-Rad, Hercules, CA). Dilute 1 part of dye concentrate to 4 parts of ddH2 O and store in a plastic bottle. The diluted solution is good for up to 2 weeks at room temperature. 2. Bovine serum albumin (BSA) protein standard (Pierce, Rockford, IL). Samples are stored as stocks of 2 mg/mL in ampules at 4 °C until use. 3. Purified PDFs are stored in 33% glycerol (v/v) by quick freeze in liquid nitrogen and stored at –80 °C (see Note 2).

2.4. PDF Substrates 1. f-Met-Ala-Ser (f-MAS, Bachem) is dissolved in ddH2 O to make 10–100 mM stock solutions. To determine the concentration of f-MAS (or other N-formylmethionyl peptides) stock, the stock solution is diluted 50–100-fold into 1 mL of 50 mM sodium phosphate, pH 7.0. Five to 10 μg of Co-EcPDF (1 μL) are added, and the reaction is allowed to proceed at room temperature for 30 min to ensure the complete release of the formate. Next, 5–10 U of FDH and 5 mM of NAD+ (final concentration) are added, and the mixture is incubated at room temperature for 30 min. The absorbance at 340 nm is measured on a UV-Vis spectrophotometer. The formate concentration is calculated using a molar absorptivity (340 nm ) of 6300 M−1 cm−1 . 2. f-ML-pNA (Bachem) stock solution is prepared by dissolving 5–10 mg of solid in 5 mL ethanol. To quantitate the f-ML-pNA stock concentration, dilute the stock (50–100-fold) into 1 mL of 1 M NaOH. Heat the solution at 95 ºC for 10 min to ensure complete hydrolysis. The solution will turn yellowish as hydrolysis of f-ML-pNA releases the chromogenic pNA. The absorbance at 405 nm is measured, and the concentration of f-ML-pNA is calculated (pNA molar absorptivity 405 nm = 10,600 M−1 cm−1 ). The stock solution (5–10 mM) is stored at -20 ºC (see Note 3).

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3. f-MK-AMC is not yet commercially available but can be easily prepared in a chemical or biochemical laboratory (16). A stock solution is prepared in DMSO and stored at -20 ºC. The concentration is determined by diluting the stock solution 10–100fold in 1 mL containing 50 mM HEPES, pH 7.0, 150 mM NaCl, and 5–10 μg of Co-EcPDF. The reaction mixture is incubated at room temperature and covered with aluminum foil for 3 h to allow the complete removal of the N-formyl moiety. Next, 5 mM DTT (final concentration) and 1 U of DPPI are added and incubated at room temperature for 1 h to allow the complete release of the chromogenic 7-amino-4methylcoumarin (AMC). The absorbance at 360 nm is measured, and the concentration of f-MK-AMC is calculated (AMC molar absorptivity e360 nm = 17,000 M−1 cm−1 ).

2.5. PDF Coupling Enzymes 1. Candida boidinii FDH (Sigma, St. Louis, MO) stock solution (5 U/μL) is prepared in 50 mM sodium phosphate, pH 7.0, and aliquots are stored frozen at –80°C (see Note 4). 2. Aeromonas proteolytica AAP (Sigma, St. Louis, MO) stock solution (50 U/mL) is prepared in 50 mM HEPES, pH 7.0, 150 mM NaCl. Aliquots are stored frozen at –20°C (see Note 5). 3. DPPI (Sigma, St. Louis, MO) stock solution (5 U/mL) is prepared in 50 mM HEPES, pH 7.0, 10 mM NaCl, 5 mM DTT. Aliquots are stored frozen at –80 °C (see Note 6).

3. Methods Since each PDF assay method has its advantages and disadvantages, we make the following recommendations. For routine kinetic assays of wild-type and mutant PDF in the absence of PDF inhibitors, the AAP-coupled assay is the best choice. f-ML-pNA is an excellent substrate of PDF, and the released p-nitroaniline has a large extinction coefficient, making this method highly sensitive. Also, the released ML-pNA is rapidly hydrolyzed by AAP, so the assay reaction can be conveniently performed in a continuous fashion. AAP is a very stable enzyme and easy to purify to near homogeneity. The disadvantage of this method is that AAP is also a metallopeptidase and therefore often strongly inhibited by PDF inhibitors, which are usually metal chelators. Both FDH and DPPI assays may be employed to screen PDF inhibitors. The FDH assay has the advantage that PDF inhibitors seldom affect the FDH reaction. However, FDH is a rather inefficient enzyme, and the coupled assay is typically performed in an end-point (discontinuous) fashion. If kinetic characterization of a PDF inhibitor is desired or if the PDF inhibitor is exceptionally potent (low KI -value), the DPPI assay is the method of choice. This method is extremely sensitive because f-MK-AMC is a very efficient substrate of PDF, MK-AMC

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is an excellent substrate of DPPI, and detection is based on fluorescence. Since DPPI, a cysteine protease, is not inhibited by PDF inhibitors, the coupled assay can be carried out in a continuous fashion in the presence of PDF inhibitors. However, for applications that require high substrate concentrations, which cause internal fluorescence quenching, the DPPI assay is best performed in the absorbance mode. 3.1. Purification of PDF 3.1.1. Bacterial Cell Growth 1. Prepare minimal media for bacterial culture. Add 10 mL of 100X nutrient mix, 1 mL of 1000X trace metal supplement, and 1 mL of 75 mg/mL ampicillin to 1 L of minimal media. 2. Inoculate 100 mL of the above minimal media with a single colony of E. coli BL21(DE3) cells harboring the pET22b-PDF expression vector and incubate overnight at 37 °C with shaking. 3. Next morning, dilute 10 mL of the overnight culture into 1 L of the prepared minimal media. Incubate at 37 °C with shaking until OD600 nm reaches 0.5–0.7. This typically takes ∼3 h. 4. Add 1 mL of 100 mM IPTG and 1 mL of 100 mM CoCl2 (or FeCl3 , or 50 mM NiCl2 ) to each flask containing 1 L of cell culture. 5. Reduce the temperature to 30 °C, and continue to shake for an additional 6 h. 6. Pellet the cells by centrifugation at 5000X g in a GS3 rotor for 5 min. 7. Wash the cell pellet with 50 mM sodium phosphate, pH 7.0. 8. The cells may be immediately used in protein purification or stored at –80 ºC until later use.

3.1.2. Purification of PDF 3.1.2.1. Co(II)-PDF-HT 1. 2. 3. 4. 5. 6. 7. 8.

If the cell pellet is frozen, slowly thaw it out on ice. For cells derived from 3 L of cell culture, add 100 mL of lysis buffer. Stir the cells in a cold room for 30 min to make a fine suspension. Lyse the cells by sonication (5 × 8 s pulses) over a 30-min period. Be sure not to heat up the solution. Remove cell debris by centrifugation at 15,000 rpm (27,000X g) (SS-34 rotor) for 30 min. Load the supernatant onto a Talon column (20 mL of resin) that has been preequilibrated in buffer B. Wash the column with the same buffer (3 × 50 mL). Elute the bound PDF protein with buffer C. Collect PDF-active fractions, which can be located by SDS-PAGE analysis or activity assays. For Co(II)-PDF, protein fractions are lightly pink colored. Protein

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purified by this method is typically free of major impurities and may be used directly in PDF activity assays. 9. Dialyze the protein against 2 L of dialysis buffer at 4 ºC (two buffer changes) to remove imidazole from the sample. If necessary, the dialyzed protein solution can be concentrated by ultrafiltration using aYM-3 membrane filter. Glycerol is added to the protein to a final concentration of 33%, and the resulting mixture is quickly frozen in liquid N2 and stored at -80 ºC. Protein concentration is determined by the Bradford method using bovine serum albumin as standard.

3.1.2.2. Ni(II)-PDF-HT 1. The cell pellet from 3 L of culture is resuspended in 100 mL of lysis buffer plus 5 mM NiCl2 . 2. Cell lysis is carried out similarly as described above. 3. The crude lysate (supernatant) is loaded onto an Ni-NTA column (20 mL of resin). Wash the column with 3 × 50 mL of buffer E. 4. Elute the bound Ni(II)-PDF-HT as described above with buffer F. 5. Pool the PDF fractions (∼50 mL) and precipitate the protein by the addition of ammonium sulfate to 80% saturation. Dissolve the precipitate in 10 mL of buffer J and load it onto a FPLC MonoQ HR 5/5 column (AKTA, Pharmacia) that has been equilibrated in buffer J. Elute the bound PDF by a linear gradient of 10 mM–1 M NaCl in buffer J. 6. Collect fractions containing significant PDF activity and analyze their purity on a 15% SDS-PAGE gel. 7. Pool the pure PDF fractions and dialyze the protein against buffer D plus 50 μM NiCl2 at 4 ºC (2 buffer changes). The resulting protein is concentrated and stored as described above.

3.1.2.3. Fe(II)-PDFH6 1. All buffers must be degassed by first applying a vacuum and then sparging with a dry inert gas (He or Ar) for 30 min. 2. Add a freshly prepared TCEP solution to the lysis buffer to give a final concentration of 1 mM. Buffers for all subsequent purification steps must contain 0.5 mM TCEP, and purification steps are carried out as fast as possible. 3. Cell lysis and purification on Talon column are carried out as described above. 4. The PDF fractions from the Talon column are concentrated by ammonium sulfate precipitation (80% saturation). 5. Dissolve the protein pellet in 5 mL of degassed 20 mM HEPES, pH 7.0, 150 mM NaCl, 1 mM TCEP. The sample is frozen and stored at –80 ºC.

3.2. FDH Coupled Assay This method utilizes the activity of Candida boidinii FDH. The formate released by PDF is oxidized into CO2 by FDH while reducing NAD+ to NADH

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(Fig. 1a). The formation of NADH is monitored at 340 nm (340 nm = 6300 M−1 cm−1 ) on a UV-Vis spectrophotometer. Peptide f-MAS is usually used as substrate, although any N-formylmethionyl peptide will also work. Co(II)substituted E. coli PDF (Co-EcPDF) has a kcat /KM -value of 2.9 × 104 M−1 s−1 toward this substrate (18). A major advantage of this method is that PDF inhibitors do not typically inhibit FDH. However, FDH is inhibited by a high salt concentration (e.g., >50 mM NaCl). In addition, FDH has low specific activity for formate (kcat of 0.9 s−1 and KM of 2.5 mM), making the FDH reaction partially rate-limiting as manifested by an early lag phase (0–20 s). Consequently, screening for PDF inhibitors by this method is often carried out in an end-point fashion. 3.2.1. Continuous Assay 1. Prepare f-MAS substrate stock solution in ddH2 O (typically, 10–100 mM). 2. To a quartz cuvette with a 1 cm optical path, sequentially add 20 μL of 10X FDH assay buffer, 20 μL of 100 mM NAD+ stock solution, 20 μL of f-MAS (1 mM final concentration), 1.0 U of FDH, a proper concentration of PDF inhibitor, and ddH2 O to adjust the total volume to 190 μL. 3. Initiate the reaction by the addition of 10 μL of diluted PDF working stock (0.1– 10 μg). 4. Monitor the reaction at 340 nm on a UV-Vis spectrophotometer.

3.2.2. End-Point Assay 1. To a microcentrifuge tube, add 30 μL of 10X FDH assay buffer, 30 μL of f-MAS (1 mM final concentration), a proper concentration of PDF inhibitor, and ddH2 O to make a total volume of 290 μL. 2. Add 10 μL of PDF (0.1–10 μg) to initiate the reaction. Allow the reaction to proceed for 60 s at room temperature (see Note 7). 3. Quench the reaction by heating in a boiling water bath for 5 min (see Note 8). 4. Cool the mixture to room temperature, and centrifuge at 14,000 rpm (17,000X g) for 5 min in a microcentrifuge. 5. Add 50 μL of 100 mM NAD+ , 0.2 U of FDH, and ddH2 O to bring it to total volume of 500 μL. 6. Allow the reaction to proceed to completion at room temperature (∼10 min). 7. Measure the OD340 nm in a UV-Vis spectrophotometer. 8. Calculate the inhibition constants.

3.3. AAP Coupled Assay This assay utilizes AAP as the coupling enzyme and f-ML-pNA as substrate (Fig. 1b). Removal of the N-formyl group by PDF produces ML-pNA, which is

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subsequently hydrolyzed by AAP into Met, Leu, and p-nitroaniline. f-ML-pNA is an excellent substrate of PDF (e.g., Co-EcPDF has a KM -value of 20 μM, a kcat -value of 38 s−1 , and a kcat /KM -value of 1.9 × 106 M−1 s−1 ). ML-pNA is also a very efficient substrate of AAP. For example, the last step, the hydrolysis of Leu-pNA, is extremely fast, with a KM -value of 38 μM, a kcat -value of 28 s−1 , and a kcat /KM -value of 7.5 × 105 M−1 s−1 . The released chromophore (pNA) has a large molar absorptivity (405 nm = 10,600 M−1 cm−1 ). These features make the AAP assay highly sensitive, convenient to perform, and relatively inexpensive. However, AAP enzyme is also a zinc metallopeptidase and may be inhibited by PDF inhibitors. Therefore, this assay is most useful for mechanistic studies of wild-type and mutant PDF in the absence of inhibitors, or screening of PDF inhibitors in the end-point fashion (15). 3.3.1. Continuous Assay 1. The assay reaction is most conveniently carried out in a disposable 1.5-mL polystyrene cuvette. 2. To the cuvette, add 100 μL of 10X AAP assay buffer, 0–200 μM f-ML-pNA, 0.8 U AAP, and ddH2 O to make a total volume of 990 μL. 3. Initiate the reaction by the addition of 10 μL of PDF (0.1–1 μg) diluted in buffer K (50 mM HEPES, pH 7.0, 100 μg/mL bovine serum albumin). 4. Monitor the reaction progress at 405 nm on a UV-Vis spectrophotometer. The initial reaction rate is calculated from the first 60 s of the reaction progress curve. 5. Reactions at the lowest and highest substrate concentrations are generally repeated with a doubled amount of AAP (1.6 U) to ensure that the PDF reaction is ratelimiting in the coupled reaction sequence.

3.3.2. End-Point Assay 1. Assay reactions are typically carried out in 1.7-mL microcentrifuge tubes with a 200-μL total reaction volume (see Note 8). 2. Add 20 μL of 10X AAP assay buffer, 0–200 μM f-ML-pNA, a proper concentration of PDF inhibitor, and ddH2 O to make a total volume of 190 μL. 3. Initiate the reaction by the addition of 10 μL of diluted PDF (0–90 ng), and incubate at room temperature for 60 s. 4. Quench the reaction by quickly placing the reaction tubes in a 95 °C water bath for 5 min. PDF inactivation typically occurs in less than 30 s. 5. Remove the tubes from the water bath, cool to room temperature, and spin for 5 min in a microcentrifuge. 6. Add 0.4 U AAP and incubate the mixture at room temperature for 30 min. 7. Transfer the reaction contents into a microcuvette and measure the absorbance at 405 nm on a UV-Vis spectrophotometer. For samples at low-substrate and/or -inhibitor concentrations, the percentage of substrate-to-product conversion should

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be calculated. This is done by comparing the amount of pNA formed and the initial concentration of f-ML-pNA. It is imperative that the percentage does not exceed 20%.

3.4. DPPI Coupled Assay The DPPI coupled assay uses N-formyl-Met-Lys-AMC as a highly efficient PDF substrate, which has a KM -value of 40 μM, a kcat -value of 26 s−1 , and a kcat /KM -value of 6.5 × 105 M−1 s−1 toward Co-EcPDF. Removal of the Nformyl moiety by PDF gives MK-AMC as a product, which is rapidly converted into dipeptide Met-Lys and 7-amino-4-methylcoumarin (KM = 2.8 μM, kcat = 5.7 s−1 , and kcat /KM = 2.1 × 106 M−1 s−1 ) (Fig. 1c). Coupled with fluorescence detection, this method is therefore extremely sensitive. This method is ideally suited for PDF inhibitor screening. PDF inhibition assays can be carried out in a continuous fashion by simply adding the proper PDF inhibitor into the reaction mixture. This latter feature is especially useful for realtime kinetic experiments and has been applied to determine the slow-binding kinetics of macrocyclic PDF inhibitors (16,20). A minor drawback of this method is the presence of slight background hydrolysis of f-MK-AMC by DPPI (KM = 82 μM, kcat = 0.06 s−1 , and kcat /KM = 760 M−1 s −1 ). This is not a major problem for routine kinetic assays or inhibitor screening involving the wild-type PDF. However, this background reaction may become significant and thus complicate the assay results involving catalytically impaired PDF mutants. 1. The following procedure is based on detection in the absorbance mode (see Note 9). Assay reactions are carried out in quartz microcuvettes. 2. Prior to use, DPPI is treated with 5 mM DTT for 30 min on ice in 50 mM HEPES, pH 7.0, 10 mM NaCl. 3. Prepare fresh 10X DPPI assay buffer each day by adding 50 mM DTT to the buffer. 4. Prepare inhibitor stock solutions (1 mM) in DMSO since many PDF inhibitors have limited solubility in aqueous solution. Working stock solutions of 10–100 μM are prepared from this original stock by diluting in DMSO. 5. A typical assay reaction has a 200-μL total volume. To a microcuvette, add 20 μL of the 10X DPPI assay buffer, 1–5 μL of working inhibitor stock solution (0–200 nM final concentration), 0.1 U of DPPI, 100 μM f-MK-AMC, and ddH2 O to bring the reaction volume to 195 μL. Place the cuvette in the spectrophotometer and monitor the reaction for 30 s at 360 nm. The slope of the curve gives the amount of background hydrolysis of substrate by DPPI. 6. The PDF reaction is then initiated by the addition of 5 μL of PDF (4.0 nM final concentration). The reaction is monitored continuously for another 60–120 s. The

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initial rate is calculated from the early region of the progress curve (0–30 s after the addition of PDF) and corrected by subtracting the background rate from above. 7. The inhibition constant (KI ) is calculated according to the equation V = Vmax × S / KM 1 + I/KI + S 

4. Notes 1. Expression of eukaryotic PDF in E. coli may suffer from low yields due to codon usage bias. This problem can be alleviated by the use of the E. coli BL21 (DE3) Rosetta strain (Novagen). This particular strain carries rare tRNA synthetases, which recognize rare codons that are normally found in E. coli but frequently encountered in G+C-rich DNA. 2. Fe(II)-PDF is very unstable when stored as a solution, even when every effort is made to remove reactive oxygen species. However, it is stable for at least 1 year when stored frozen at –80 °C. We recommend that it be stored in small aliquots at –80 °C and that a fresh aliquot be thawed on ice and used at least once a day. Co(II)-, Ni(II)-, and Zn(II)-EcPDF are more stable and can be stored at 4 °C for weeks without significant loss of activity. 3. f-ML-pNA has a maximal solubility of ∼200 μM in aqueous solution. Prepare the solution at neutral pH. Acidic pH facilitates hydrolysis of the N-formyl group, whereas basic pH promotes hydrolysis of the pNA group. 4. Formate dehydrogenase is sensitive to chloride ions. Samples come as lyophilized powder. One unit will oxidize 1.0 μmole of formate to CO2 per min in the presence of -NAD at pH 7.6 at 37 °C. It is less problematic when the assay is monitored at 365 nm. Co(II)-PDF has strong absorption near 340 nm. 5. AAP is a zinc-containing enzyme. Samples come in lyophilized powder. Often, AAP obtained from Sigma contains background endopeptidase activity. The Prescott and Wilkes method employing the gelatin digestion assay can measure the endopeptidase activity (21). The residual enzyme activity can often be removed by heating the stock enzyme solution at 65 °C for 2 h. One unit will hydrolyze 1.0 μmole of L-leucine-p-nitroanilide to L-leucine and p-nitroaniline per min at pH 8.0 at 25 °C. Working stock samples can be stored at 4 °C for up to 2 weeks. Avoid freezethaw cycles. Since this assay is the method of choice in our lab for routine PDF characterization, we found it to be economical to purify AAP from Aeromonas proteolytica by the Prescott and Wilkes method (21). 6. Dipeptidyl peptidase I from bovine spleen is a cysteine protease and must be reactivated with 5 mM dithiothreitol (DTT) for 30 min on ice in 50 mM HEPES, pH 7.0, 10 mM NaCl before use. Samples come in lyophilized powder. The enzyme is very unstable and must be stored frozen. One unit will produce 1 μmole of

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Gly-Phe-NHOH from Gly-Phe-NH2 and hydroxylamine per min at pH 6.8 at 37 °C using DL-phenylalanine hydroxamic acid as the standard. 7. The reaction time should be short due to rapid inactivation of Fe-PDF under the assay conditions. For Co- and Ni-PDF, which are stable, smaller amounts of PDF and longer reaction times (5–10 min) are recommended. 8. Smaller sample volumes are easier to quench with heating. Alternatively, the reaction can be terminated by the addition of 25 μL of 1 M HCl. The reaction is neutralized by the addition of 25 μL of 2 M HEPES, pH 7.0. Make sure that the final reaction is at pH ∼7 by testing it with a pH paper before adding AAP. If the pH is not ∼7, adjust the pH to 7 by adding additional 2 M HEPES, pH 7.0 solution. 9. PDF inhibition assay can also be monitored in the fluorescence mode on an AmincoBowman Series 2 Luminescence spectrometer and adapts a similar procedure to the absorbance mode. The excitation and emission wavelengths are set at 380 and 460 nm, respectively.

Acknowledgments This work was supported by NIH Grant AI40575. References 1 Giglione, C., Pierre, M., and Meinnel, T. (2000) Peptide deformylase as a target 1. for new generation, broad spectrum antimicrobial agents. Mol. Microbiol. 36, 1197–1205. 2 Pei, D. (2001) Peptide deformylase: A target for novel antibiotics? Emerging Ther. 2. Targets 5, 23–40. 3 Yuan, Z, Trias, J., and White, R. J. (2001) Deformylase as a novel antibacterial 3. target. Drug Discovery Today 6, 954–961. 4 Clements, J. M., Beckett, R. P., Brown, A., Catlin, G., Lobell, M., Palan, S., 4. Thomas, W., Whittaker, M., Wood, S., Salama, S., Baker, P. J., Rodgers, H. F., Barynin, V., Rice, D. W., and Hunter, M. G. (2001) Antibiotic activity and characterization of BB-3497, a novel peptide deformylase inhibitor. Antimicrob. Agents Chemother. 45, 563–570. 5 Meinnel, T., Mechulam, Y., and Blanquet, S. (1993) Methionine as translation start 5. signal: A review of the enzymes of the pathway in Escherichia coli. Biochimie 75, 1061–1075. 6 Ben-Bassart, A., Baur, K., Chang, S. Y., Myambo, K., Boosman, A., and Chang, S. 6. (1987) Processing of the initiation methionine from proteins: Properties of the Escherichia coli methionine aminopeptidase and its gene structure. J. Bacteriol. 169, 751–757. 7 Mazel, D., Pochet, S., and Marliere, P. (1994) Genetic characterization of 7. polypeptide deformylase, a distinctive enzyme of eubacterial translation. EMBO J. 13, 914–923.

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8 Meinnel, T., and Blanquet, S. (1994) Characterization of the Thermus thermophilus 8. locus encoding peptide deformylase and methionyl-tRNA(fMet) formyltransferase. J. Bacteriol. 176, 7387–7390. 9 Margolis, P. S., Hackbarth, C. J., Young, D. C., Wang, W., Chen, D., Yuan, Z., 9. White, R., Trias, J. (2001) Peptide deformylase in S. aureus: Resistance to inhibition is mediated by mutations in the formyltransferase gene. Antimicrob. Agents Chemother. 44, 825–1831. 10 Rajagopalan, P. T. R., Yu, X. C., and Pei, D. (1997) Peptide deformylase: A new 10. type of mononuclear iron protein. J. Am. Chem. Soc. 119, 12418–12419. 11 Groche, D., Becker, A., Schlichting, I., Kabsch, W., Schultz, S., and 11. Wagner, A. F. V. (1998) Isolation and crystallization of functionally competent Escherichia coli peptide deformylase forms containing either iron or nickel in the active site. Biochem. Biophys. Res. Comm. 246, 342–346. 12 Nguyen, K. T., Hu, X., Colton, C., Chakratarti, R., Zhu, M. X., and Pei, D. (2003) 12. Characterization of a human peptide deformylase: Implications for antibacterial drug design. Biochemistry 42, 9952–9958. 13 Lazennec, C., and Meinnel, T. (1997) Formate dehydrogenase-coupled spectropho13. tometric assay of peptide deformylase. Anal. Biochem. 244, 180–182. 14 Rajagopalan, P. T. R., Datta, A., and Pei, D. (1997) Purification, characterization, 14. and inhibition of peptide deformylase from Escherichia coli. Biochemistry 36, 13910–13918. 15 Wei, Y., and Pei, D. (1997) Continuous spectrophotometric assay of peptide 15. deformylase. Anal. Biochem. 250, 29–34. 16 Nguyen, K. T., Hu, X., and Pei, D. (2004) Slow-binding inhibition of peptide 16. deformylase by cyclic peptidomimetics as revealed by a new spectrophotometric assay. Bioorg. Chem. 32, 178–191. 17 Rajagopalan, P. T. R., and Pei, D. (1998) Oxygen mediated inactivation of peptide 17. deformylase. J. Biol. Chem. 273, 22305–22310. 18 Rajagopalan, P. T. R., Grimme, S., and Pei, D. (2000) Characterization of 18. Cobalt(II)-substituted peptide deformylase: Function of the metal ion and the catalytic residue Glu-133. Biochemistry 39, 779–790. 19 Hu, X., Nguyen, K. T., Jiang, V. C., Lofland, D., Moser, H. E., and Pei, D. (2004) 19. Macrocyclic inhibitors for peptide deformylase: A structure-activity relationship study of the ring size. J. Med. Chem. 47, 4941–4949. 20 Prescott, J. M., and Wilkes, S. H. (1976) Aeromonas aminopeptidase. Methods 20. Enzylmol. 45, 530–543.

11 A Method to Assay Penicillin-Binding Proteins Michael J. Pucci and Thomas J. Dougherty

Summary Key enzymes that assemble the bacterial cell wall are also the target of the -lactam class of antibiotics. The covalent binding of labeled penicillin to these proteins has been used in numerous studies in drug discovery, antibiotic mechanisms of action and resistance, and cell wall physiology. Methods to label and measure penicillin binding proteins in two prototypical organisms, a Gram-negative (Escherichia coli) and Gram-positive (Staphylococcus aureus), are described. The methods discussed include identifying penicillin-binding proteins in both intact cells (in vivo measurements) and isolated cell membranes.

Key Words: penicillin-binding proteins; peptidoglycan; membrane protein; fluorography.

1. Introduction The penicillin-binding proteins (PBPs) are a set of enzymes catalyzing terminal reactions of bacterial peptidoglycan biosynthesis (1–3). The ability to use radioactive or fluorescent labeled penicillin to identify this subset of bacterial proteins is the genesis of the terminology. In fact, these proteins, which are situated on the outer surface of the bacterial cytoplasmic membrane, possess key cell wall assembly enzymatic activities, which include transglycosylation and transpeptidation, along with endopeptidase and carboxypeptidase functions. The transpeptidation and carboxypeptidase functions are inhibited when these proteins are acylated by a -lactam antibiotic, with the -lactam ring opening and forming a covalent bond with an active-site serine residue found in a highly conserved binding pocket present in PBPs (4). Much of our initial knowledge about the roles of individual PBPs came from work done primarily in the From: Methods in Molecular Medicine, Vol. 142: New Antibiotic Targets Edited by: W. Scott Champney © Humana Press Inc., Totowa, NJ

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Gram-negative rod-shaped organism E. coli and the Gram-positive rod Bacillus subtilis (5–7). These proteins were found to play roles in both peptidoglycan synthesis and cell morphogenesis. However, much remains to be understood concerning their specific roles in cell wall growth and cell division. Working with E. coli, Spratt published an indirect estimate of the numbers of individual PBPs in an average E. coli cell (8). Nearly two decades later, Dougherty et al. directly calculated the numbers of molecules per cell of individual PBPs by measuring both the cell numbers present and the radioactivity in the individual PBPs from a precise quantity of cells (9). Pucci and Dougherty published a similar analysis in Staphylococcus aureus in 2002 (10). The results of these studies indicated that the numbers of the higher–molecularweight PBPs are relatively low, as few as 120 per cell, in both of these species. Penicillin-binding proteins are the targets for -lactams, one of the most effective and widely used classes of antibiotics for the treatment of bacterial infections (11). These include the penicillins and cephalosporins, which have saved innumerable lives for more than 60 years. Many key species of bacterial pathogens have been found to possess modified PBPs that exhibit greatly reduced affinities for -lactam antibiotics, leading to increased levels of antibiotic resistance. The reduced-affinity PBPs have been shown in many cases to be the result of promiscuous genetic exchange among bacteria (1). PBP assays have been used in competition assays with labeled penicillin to obtain PBP binding affinity data for individual -lactam compounds, as well as the kinetics of binding in the study of PBP–-lactam interactions. This has resulted in the discovery and characterization of many new and improved antibacterial agents in this class. We describe in this chapter methods to assay PBPs in representative Gram-negative and Gram-positive bacterial species. 2. Materials 2.1. Cell Culture, Lysis, and Membrane Preparation 1. Tryptone broth and Brain-Heart-Infusion broth (Difco, Detroit, MI). Lennox L broth or LB (Gibco BRL, Gaithersburg, MD) (see Note 1). 2. Agitation on a gyratory shaking incubator from New Brunswick Scientific Co. For E. coli and staphylococci, vigorous shaking at ∼250 rpm is used (see Note 2). 3. Bacterial growth is monitored using a Spectronic 20 spectrophotometer or equivalent at 600 nm. 4. Sonication buffer: 0.05 M Tris-HCl, pH 7.5, containing 1 mM MgCl2 , 1 mM 2-mercaptoethanol, and 5 μg/ml DNase. 5. Proteins are assayed by either the bicinchoninic acid assay (BCA; Pierce) or Lowry assay (12).

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6. Staphylococcal lysis buffer: 50 mM Tris-HCl, pH 7.0, 10 mM MgCl2 , 2.5 μg of DNase/mL, and 2.5 μg of RNase/mL followed by addition of lysostaphin (Sigma) to a final concentration of 200 μg/mL. 7. The Bead-Beater was purchased from Bio-Spec Products (Bartlesville, OK). Glass beads are obtained from this same vendor. The sonicator, Vibra-Cell, Model VC600, was purchased from Sonics and Materials (Danbury, CT) (see Note 3).

2.2. SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE) 1. Separating gel: 10 mL of 1.5 M Tris-HCl buffer pH 8.8, 16.6 mL of Bis-acrylamide (0.2:24 v/v), 0.4 mL of 10% SDS, and 13 mL of water (see Note 4) for a total volume of 40 mL. 2. Stacking gel: 2.5 mL of 0.5 M Tris-HCl buffer pH 6.8, 1.7 mL of Bis-acrylamide (0.2:24 v/v), 0.1 mL of 10% SDS, and 5.6 mL of water for a total volume of 9.9 mL. 3. Running buffer, pH 8.3, 10X stock: Tris base 15 g/L, glycine 72 g/L, and SDS 5 g/L. 4. Sample buffer, 2X: 2.5 mL of 0.5 M Tris-HCl, pH 6.8, 2.0 mL of 10% SDS, 2.0 mL glycerol, 3.3 mL of water, and 0.1 mL of 2-mercaptoethanol. 5. Staining solution (1 L): 500 mL of methanol, 100 mL of glacial acetic acid, 400 mL of water, and 2.75 g of Coomassie Blue. 6. Destaining solution (1 L): 100 mL of methanol, 100 mL of glacial acetic acid, and 800 mL of water.

3. Methods PBPs are usually assayed by an indirect method using labeled -lactam antibiotics, most often radiolabeled penicillin. This material must be customsynthesized, as it is generally not available commercially. The [3 H] version of labeled penicillin is made as an ethylpiperidinium salt, which is soluble in both acetone and water. The material is stored in acetone as small volume aliquots (40–60 μL, based on specific activity) in 1.5 mL micro fuge tubes at –80°C and is stable for up to 2 years. Just prior to use, the acetone is removed with a gentle stream of nitrogen, and the dried residue in the tube is resuspended with an appropriate volume of phosphate buffer. Alternatively, a fluorescent penicillin derivative (Bocillin), which is currently available commercially, may be used. The labeling of the PBPs can either be in vivo using whole cells or by using isolated membrane proteins. Labeled PBPs then can be separated on SDS-PAGE gels where band migration distances (and thus apparent molecular weights) and relative amounts of labeled antibiotic can be analyzed. The utilization of isolated membrane proteins is the more common method often resulting in gels with better appearances than those from in vivo labeling experiments. However, accurate quantitation in regard to the amount of input cells is very difficult using

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membrane proteins because of variable losses during the membrane preparation process. Also, membrane preparation methods differ among bacterial genera. Examples of PBP assays from both a Gram-positive and Gram-negative bacteria using both in vivo and isolated membrane protein methods are discussed in this chapter (see Note 5). 3.1. Culture Growth Conditions—E. coli 1. A single bacterial colony is inoculated into 5 mL of tryptone broth and grown overnight at 30°C with agitation. 2. For in vivo labeling, the next morning, a 200-fold dilution of the overnight culture is made in 40 mL of LB medium in a flask and grown at 35°C with agitation (∼250 rpm) in a shaking incubator. 3. For membrane preparations, the next morning, a 200-fold dilution of the overnight culture is made in 500 mL of LB medium in a flask and grown at 35°C with agitation (∼250 rpm) in a shaking incubator. 4. Growth is monitored in a spectrophotometer at 600 nm. Experiments are started in the A600 range of 0.3 to 0.4, which is the mid-logarithmic growth phase and represents about 2 × 108 cells per mL.

3.2. In vivo Labeling of PBPs—E. coli 1. Exactly 1 mL of mid-logarithmic phase cells is collected and centrifuged in microfuge tubes for 2 min at 12,000X g. 2. The supernatant is removed, and the cells are resuspended in 50 μL of 50 mM sodium phosphate buffer, pH 7.0. 3. The samples are subjected to three cycles of freeze-thaw from –80°C to 37°C, for 3 min at each temperature. 4. After the last thaw, 2 μg of radiolabeled penicillin ([3 H]benzylpenicillin, ethylpiperidium salt; 27.2 Ci/mmol) are added immediately, for a final concentration of 40 μg/mL. 5. The samples are incubated at 37°C for 20 min. 6. The reaction is terminated by addition of 25 μL of 20% Sarkosyl detergent. 7. To prepare samples for SDS-PAGE, 25 μL of SDS sample buffer containing 2-mercaptoethanol are added followed by incubation at 100°C for 5 min.

3.3. Membrane Preparations: Sonication and Differential Centrifugation—E. coli 1. E. coli cells are grown in LB medium in culture volumes of 500 mL at 35°C with agitation. 2. When the cells reach an A600 of 0.4 to 0.5, they are harvested by centrifugation at 7,000X g for 10 min at 4°C, washed once with 0.05 M of potassium phosphate buffer, followed by resuspension in 2 volumes of sonication buffer.

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3. The cells are then sonicated with cooling in an ice bath until 95% or greater breakage occurs. This may require several cycles with cooling in between. 4. Unbroken cells and cellular debris are removed by a low-speed centrifugation at 3,000X g. Retain the sonicated supernatants. 5. Supernatent fluids from the low-speed centrifugation are centrifuged to pellet membranes by ultracentrifugation at 100,000X g for 30 min at 4°C. Membranes are resuspended in sodium phosphate buffer, pH 7.0, and washed once at 100,000X g at 4°C for 30 min. Pellets are resuspended in 500 μL of phosphate buffer and stored at –70°C if not used immediately. 6. Small samples (5 μL) of the pellet resuspension are taken for protein concentration determinations by the bicinchoninic acid assay or Lowry assay, and 100 μg of protein were used per gel lane.

3.4. Membrane Preparations: Mechanical Breakage—E. coli 1. E. coli cells are grown in LB medium in culture volumes of 500 mL at 35°C with agitation. 2. When the cells reach an A600 of 0.4 to 0.5, they are harvested by centrifugation at 7,000X g for 10 min at 4°C, 3. The cells are resuspended in 10 mL of sodium phosphate buffer, pH 7.0, and broken with a Bead-Beater. 4. Glass beads (35 g) are added, and the mixture is subjected to eight cycles of agitation (15 s) in an ice-jacketed chamber. 5. Beads are separated from broken cells by filtration over a sintered glass filter, and the membranes are collected by ultracentrifugation at 155,000X g for 30 min at 4°C. 6. Membranes are resuspended and washed once with sodium phosphate buffer at 4°C, again pelleted by ultracentrifugation at 155,000X g, and resuspended in a final volume of 250 μL in sodium phosphate buffer. If membranes are not to be used immediately, pellets resuspended in a small volume of phosphate buffer can be stored frozen at –70°C. 7. Samples are taken for protein concentration determinations by the bicinchoninic acid assay or Lowry assay, and 100–200 μg of protein are used per gel lane.

3.5. Culture Growth Conditions—Staphylococcus aureus 1. A single bacterial colony is inoculated into 5 mL of BHI broth and grown overnight at 35°C with agitation. 2. For in vivo labeling, the next morning, a 200-fold dilution of the overnight culture is made in 40 mL of BHI medium in a flask and grown at 35°C with agitation (∼250 rpm) in a shaking incubator. 3. For membrane preparations, the next morning, a 200-fold dilution of the overnight culture is made in 1 L of BHI medium in a flask and grown at 35°C with agitation (∼250 rpm) in a shaking incubator.

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4. Growth is monitored in a spectrophotometer at 600 nm. Experiments are started in the A600 range of 0.3 to 0.4, which is the mid-logarithmic growth phase and represents about 2 × 108 cells per mL.

3.6. In vivo Labeling of PBPs—S. aureus 1. One-mL samples of cells from mid-logarithmic phase cultures of S. aureus are immediately centrifuged and resuspended in 50 μL of staphylococcal lysis buffer. 2. Two microliters of radiolabeled penicillin ([3 H]benzylpenicillin, ethylpiperidium salt, 26.5 Ci/mmol; final concentration, 40 μg/mL) are immediately added, followed by a 20-min incubation at 37°C (see Note 6). 3. Five microliters of lysostaphin are added to a final concentration of 200 μg/mL, and samples were incubated at 37°C for an additional 5 min to allow cell lysis to occur, as indicated by clearing of the sample solution. 4. Immediately after lysis occurs, an SDS-PAGE sample buffer is added and the samples are boiled for 5 min. 5. The entire sample volumes are loaded into lanes on an SDS-PAGE gel and subjected to electrophoresis. 6. Following electrophoresis, gels are subjected to fluorography to reveal the positions of the radiolabeled PBPs. Bands on films can be quantitated and analyzed.

3.7. Membrane Preparations: Mechanical Breakage—S. aureus 1. S. aureus strains are grown to mid-logarithmic phase in BHI broth with aeration. Volumes of 500 or 1000 mL will usually provide sufficient cell numbers to prepare membranes for a reasonable number of experiments. 2. When the cells reach an A600 of approximately 0.6, they are harvested by centrifugation at 7,000X g for 10 min at 4°C. The cells are washed once and resuspended in 10 mL of ice-cold 10 mM sodium phosphate buffer, pH 7.0. 3. Alternatively, pellets can be frozen at –70°C overnight and thawed and resuspended prior to further use. 4. Glass beads (∼35 g) of 0.1 mm diameter are added, and the mixture is subjected to 8 cycles of agitation (20 s) in a Bead-Beater using an ice-jacketed chamber allowing 5 min of cooling in between bursts. 5. The lysates are passed through a coarse-grade sintered glass filter to remove the beads. The filtrate then is centrifuged at 3000 rpm to remove any remaining glass beads and unbroken cells. 6. The supernatant is centrifuged at 100,000X g for 30 min at 4°C to pellet membranes. The membranes are resuspended in sodium phosphate buffer and washed once by centrifugation at 100,000X g. 7. Membranes are resuspended to a final volume of 1 mL in 10 mM sodium phosphate buffer, pH 7.0. Remove a small amount, ∼10 μL, for protein determination. If membranes are not to be used immediately, the resuspended pellets can be stored

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frozen at –70°C. Membranes can also be aliquotted and frozen after protein concentration is estimated. 8. Protein concentration determinations are obtained by the bicinchoninic acid assay or Lowry assay, and 100 μg of protein are used per gel lane.

3.8. SDS-Polyacrylamide Gel Electrophoresis 1. To maximize resolution of PBP bands, gels should be run on a long bed gel (≥20 cm) at a constant current of 10 mA at 4°C for 16 h. This is important, as some PBPs have very similar molecular weights and can be challenging to resolve. 2. Prepare a 1.5-mm-thick, 10% separating gel using glass plates (one with a comb notch) and plastic spacers on both sides and bottom of the sandwich. The spacers and plate sandwich are held together with alligator clips. Degas the gel solution under vacuum for 5 min. Add 160 μL of 10% ammonium persulfate and 16 μL of TEMED and swirl to mix. Pour the separating gel immediately, leaving space for the stacking gel, and overlay with water-saturated isobutanol. Allow 1 h for complete polymerization. 3. Pour off the isobutanol, and rinse the top of the gel with water. 4. Prepare the 5% stacking gel. Degas the gel solution under vacuum for 5 min. Add 100 μL of freshly made 10% ammonium persulfate and 5 μL of TEMED, and swirl to mix. Pour the stacking gel and quickly insert the comb. The stacking gel should polymerize within 30 min. 5. Prepare the running buffer by diluting 100 mL of the 10X running buffer with 900 mL of water in a beaker with stirring. 6. Once the stacking gel has set, carefully remove the comb and use a 3-mL syringe with a 22-gauge needle to wash the wells with running buffer. Remove the bottom spacer between the glass plates. 7. Attach the glass plates containing the complete gel to the gel unit with clamps, and fill the gel unit with runner buffer. Using a glass pipette with the tip bent at a 45° angle by heating, remove any air bubbles from the bottom interface of the gel, where the spacer was removed. Complete filling of the gel unit with running buffer. Check for and remove air bubbles from loading areas between the teeth of the stacking gel by flushing with buffer using a hand pipettor with a disposable sequencing gel tip. 8. Load samples with a pipettor and narrow, disposable sequencing tips into each well, including molecular-weight markers. Run gel overnight at 10 mA constant current. The dye front should be run to the bottom of the gel or off the gel completely.

3.9. Fluorography and Quantitation of PBPs 1. Following electrophoresis, the gel is fixed and stained and prepared for fluorography with En3 Hance fluor. After drying on a gel dryer, the dried gel is exposed to preflashed X-OMAT AR X-ray film (Kodak) at –70°C (see Note 7). The film is first oriented as to position on the gel by marking with a marker on both the gel

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Fig. 1. Fluorograph of E. coli K-12 isolated membranes in PBP competition experiments with a cephalosporin. PBP designations are indicated on the left. Radiolabeled penicillin is added to a final concentration of 40 μg/mL. The concentrations of competing antibiotic in lanes a to h are 0, 0.125, 0.25, 0.5, 1.0, 2.0, 4.0, and 8.0 μg/mL. and the film (see Note 8). The film is developed and fixed to reveal the positions of the radiolabeled bands (see Note 9). Examples of results for E. coli and S. aureus are shown in Figs. 1 and 2. 2. Penicillin binding can be quantified by scanning PBP fluorographs using a scanning laser densitometer or an image analyzer device to generate areas under the peaks corresponding to the various PBPs in each gel lane. The areas are plotted, and best-fit sigmoidal curves are determined by using curve-fitting software such as Prism or SigmaPlot. Kinetics of binding and IC50 values can be determined for PBPs and -lactam antibiotics. An example is shown in Fig. 3.

1 2a 2 3

a

b

c

d

e

f

g

4

Fig. 2. Fluorograph of S. aureus RN4220 isolated membranes in PBP competition experiments with methicillin. PBP designations are indicated on the left. Radiolabeled penicillin is added to a final concentration of 40 μg/mL. The concentrations of competing antibiotic in lanes a to g are 0, 8, 16, 32, 64, 128, and 0 μg/mL.

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Fig. 3. E. coli PBP binding curve for a cephalosporin antibiotic. Areas were determined by scanning laser densitometry of individual lanes on fluorographs. Areas under the peaks are plotted versus the log of the concentrations (μg/mL) of the competing antibiotic. Best-fit sigmoidal curves were plotted using GraphPad Prism software.

3.10. Detection Using a Fluorescent Penicillin Derivative 1. Substitute Bocillin FL (sodium salt-Invitrogen, Carlsbad, CA) for [3 H] or [14 C] penicillin (13,14). Bocillin FL is dissolved in water and stored in aliquots at –20°C in the dark. Bocillin is diluted and added in final concentrations of 5–10 μM and incubated with membranes in either phosphate (pH 7.4) or Tris-Cl (pH 7.5) and NaCl (0.15–0.5 M) for 30 min at 35°C. The membrane preparations are then treated as above by boiling in SDS containing sample buffer to stop the reaction. 2. Samples are loaded onto SDS-polyacrylamide gels and electrophoresed as described above. After electrophoresis, the gel is washed briefly with distilled water. 3. The PBPs with the bound fluorescent Bocillin can be visualized by eye with a UV lamp. Quantitation of the labeled PBPs can be accomplished with an instrument such as a FluorImager 595 (Molecular Dynamics, Sunnyvale, CA) by excitation at 488 nm and detection at 530 nm. Other imaging systems such as the Storm PhosphoImager (Molecular Dynamics) or Alphaimager (Alpha Innotech, San Leandro, CA) can be employed.

4. Notes 1. Suggested bacterial culture media are listed. Other media may also be appropriate, but adjustments may be necessary to adjust for varied cell numbers. 2. Agitation of cultures was done in New Brunswick shaking incubators. It is important to maintain vigorous aeration of E. coli and staphylococcal cultures to ensure optimal

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growth rates. Other Gram-positive bacteria such as streptococci and enterococci must be grown without agitation and aeration. Care must be taken when using either a sonicator or Bead-Beater to prevent cells and lysates from warming. Cooling periods in an ice-water bath are essential. All solutions should be prepared in distilled water or water with a resistivity of 18.2 M-cm. The methods should work for most strains dependent upon satisfactory growth in culture medium. Radiolabeled penicillin may require a custom synthesis from the vendor. If [3 H]benylpenicillin is used, films should be exposed for 3–7 days. If [14 C]benzylpenicillin is used, films should be exposed for 2–4 weeks. Film is preflashed as described in (15) to obtain a linear response. It is important to carefully orient the film to the gel so that the exposed bands can be aligned with the molecular-weight marker bands for size estimation. Films can be developed manually using standard X-ray film developer and fixer or by automated X-ray film processors, if available.

Acknowledgments We would like to thank Alexander Tomasz and Roberta Fontana for their assistance in helping us become proficient in the methodologies described in this work. We also thank Regine Hakenbeck and Brigitte Berger-Bachi for helpful discussions over the years. References 1 Macheboeuf, P., Contreras, C., Job, V., Dideberg, O., and Dessen, A. (2006) 1. Penicillin binding proteins: Key players in bacterial cell cycle and drug resistance processes. FEMS Microbiol. Rev. 30, 673–691. 2 Spratt, B. G. (1975) Distinct penicillin-binding proteins involved in the division, 2. elongation, and shape of Escherichia coli K-12. Proc. Natl. Acad. Sci. USA 72, 2999–3003. 3 Koch, A. L. (2000) Penicillin-binding proteins, beta-lactams, and lactamases: 3. Offenses, attacks, and defensive countermeasures. Crit. Rev. Microbiol. 26, 205–220. 4 Goffin, C., and Ghuysen, J.-M. (1998) Multimodular penicillin-binding proteins: 4. An enigmatic family of orthologs and paralogs. Microbiol. Mol. Biol. Rev. 62, 1079–1093. 5 Young, K. D. (2001) Approaching the physiological functions of penicillin-binding 5. proteins in Escherichia coli. Biochemie 83, 99–102. 6 McPherson, D. C., and Popham, D. L. (2003) Peptidoglycan synthesis in the 6. absence of class A penicillin-binding proteins in Bacillus subtilis. J. Bacteriol. 185, 1423–1431.

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7 Stewart, G. C. (2005) Taking shape: Control of bacterial cell wall biosynthesis. 7. Mol. Microbiol. 57, 1177–1181. 8 Spratt, B. G. (1977) Properties of the penicillin-binding proteins of Escherichia 8. coli K-12. Eur. J. Biochem. 72, 341–352. 9 Dougherty, T. J., Kennedy K., Kessler, R. E., and Pucci, M. J. (1996) Direct 9. quantitation of the number of individual penicillin-binding proteins per cell in Escherichia coli. J. Bacteriol. 178, 6110–6115. 10 Pucci, M. J., and Dougherty, T. J. (2002) Direct quantitation of the number 10. of individual penicillin-binding proteins per cell in Staphylococcus aureus. J. Bacteriol. 184, 588–591. 11 Tipper, D. J., and Strominger, J. L. (1965) Mechanism of action of penicillins: 11. A proposal based on their structural similarity to acyl-D-alanyl-D-alanine. Proc. Natl. Acad. Sci. USA 54, 1133–1141. 12 Lowry, O. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J. (1951) Protein 12. measurement with the Folin phenol reagent. J. Biol. Chem. 193, 265–275. 13 Zhao, G., Meier, T. I., Kahl, S. D., Gee, K. R., and Blaszczak, L. C. (1999) 13. Bocillin FL, a sensitive and commercially available reagent for detection of penicillin-binding proteins. Antimicrob. Agents Chemother. 43, 1124–1128. 14 Kaczmarek, F. S., Gootz, T. D., Dib-Hajj, F., Shang, W., Hallowell, S., and 14. Cronan, M. (2004) Genetic and molecular characterization of -lactamase-negative ampicillin-resistant Haemophilus influenzae with unusually high resistance to ampicillin. Antimicrob. Agents Chemother. 48, 1630–1639. 15 Laskey, R. A., and Mills, A. D. (1975) Quantitative film detection of 3H and 14C 15. in polyacrylamide gels by fluorography. Eur. J. Biochem. 56, 335–341.

12 A Method to Assay Inhibitors of Lipopolysaccharide Synthesis Marcy Hernick and Carol A. Fierke

Summary Treatment of Gram-negative bacterial infections is complicated by innate and acquired drug resistance resulting in a limited number of effective antibiotics. Several Gram-negative bacteria, for which current therapies are ineffective, have recently been identified as potential bioterror agents. These findings highlight the need for new antibiotics, specifically antibiotics that act on new drug targets to circumvent drug resistance. Potential targets in Gram-negative bacteria include enzymes involved in the biosynthesis of lipopolysaccharides (LPS) that form outer membranes of these organisms. UDP-3-O-(R-3-hydroxymyristoyl)-N-acetylglucosamine deacetylase (LpxC) catalyzes the committed step in the biosynthesis of the lipid A portion of LPS. Therefore, inhibitors of this enzyme have the potential to serve as antibiotics, and efforts toward the development of LpxC inhibitors are currently underway. Here we describe methods for assaying LpxC inhibitors, including methods for measuring deacetylase activity and binding affinity for LpxC, which will be useful for the development of LpxC inhibitors.

Key Words: lipopolysaccharide; LPS; LpxC; lipid A; inhibitor; metal-dependent deacetylase.

1. Introduction Lipopolysaccharides (LPS) are negatively charged molecules that form the outer membranes of Gram-negative bacteria and act as barriers to prevent entry of small molecules, thereby contributing to the viability and innate resistance of these organisms (1,2). Structurally, LPS consists of three regions: O-antigen, core, and lipid A. The O-antigen and core portions of LPS are variable oligosaccharides, while lipid A is a glucosamine phospholipid that From: Methods in Molecular Medicine, Vol. 142: New Antibiotic Targets Edited by: W. Scott Champney © Humana Press Inc., Totowa, NJ

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has relatively conserved architecture across different organisms (1). Lipid A is responsible for anchoring LPS to the outer membrane and is also the portion of LPS responsible for stimulating the immune system in Gram-negative sepsis (1–3). The loss of lipid A results in decreased viability and increased sensitivity to antibiotics; therefore, inhibitors of lipid A biosynthesis have the potential to function as antibiotics and/or anti-endotoxins (1,2,4). UDP-3-O-(R-3-hydroxymyristate)-N-acetyl-glucosamine deacetylase (LpxC) is a metal-dependent enzyme that catalyzes the committed, and second overall, step in the biosynthesis of lipid A—the conversion of UDP-3-O-(R-3hydroxymyristate)-N-acetyl-glucosamine to UDP-3-O-(R-3-hydroxymyristate)glucosamine and acetate (5,6). Consequently, LpxC is currently being pursued as a target for the development of antibiotics (4,7–12). The appeal of LpxC as a drug target is enhanced by the finding that it is a metalloenzyme (6) with known information regarding its biochemical mechanism (13–16) and a solved three dimensional structure (17,18). LpxC inhibitors typically contain a group capable of chelating the catalytic metal ion (i.e., hydroxamate, phosphonate, carboxylate) (4,7–12), based on knowledge from past successes in developing metalloenzyme inhibitors (19–23). In the future, LpxC inhibitors will be optimized for potency and specificity as more biochemical and structural data become available. Here we describe assays for in vitro evaluation of LpxC inhibitors using purified LpxC, including assays for measuring the effect of inhibitors on deacetylase activity and inhibitor binding affinity for LpxC. 2. Materials 2.1. Preparation of [14 C]-UDP-3-O-(R,S-hydroxymyristoyl) -N-acetyl-glucosamine 1. 2. 3. 4. 5. 6.

1 M bis-tris, pH 6 (Research Organics Inc., Cleveland, OH). 50 mM bis-tris, pH 6, 1 M NaCl. 10 mM bis-tris, pH 6. 0.2 M NaCl (see Note 1). Filter (0.2 μm) aqueous solutions and store at room temperature. Freshly prepared solutions: 0.4 M HEPES, pH 8 (Fisher Scientific), 10 mg/mL bovine serum albumin fatty acid-free (BSA; Sigma, St. Louis, MO), and 100 mM methyl methanethiosulfonate (MMTS; Sigma, St. Louis, MO). 7. [14 C]-UDP-N-acetyl-glucosamine (NEN Life Science Products, Boston, MA) stored at –20 °C. 8. Disposable 10 mL and 60 mL luer-lok syringes (Fisher Scientific). 9. Costar 3620 1.7 mL microcentrifuge tubes (Corning Incorporated, Corning, NY) (see Note 2).

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10. Poly prep columns, 0.8 × 4 cm (Bio-Rad, Hercules, CA) filled with 2 mL DEAEsepharose fast flow resin (GE Healthcare BioSciences, Piscataway, NJ). Prior to initiating reactions: Wash column with 18 mL H2 O (to remove ethanol), equilibrate column with 16 mL 1 M bis-tris, pH 6, followed by 8 mL 50 mM bis-tris, pH 6, 1 M NaCl, and finally 8 mL 10 mM bis-tris, pH 6. Use a 60 mL disposable syringe fitted with column adapter to apply pressure (air) to speed the flow of solutions through the column. 11. C18 sep-pak (Waters, Milford, MA) washed with 10 mL methanol followed by 20 mL H2 O. Attach equilibrated sep-pak onto bottom of DEAE-sepharose column. 12. R S-3-hydroxy-myristate-ACP (2-3 mM) and LpxA (11–39 mg/mL) stored at –80 °C (see Note 3).

2.2. LpxC Deacetylase Assay 1. Costar 3620 1.7 mL microcentrifuge tubes, Costar 4826 (0.1–10 μL) tips, and Costar 4863 (1–200 μL) tips (Corning Incorporated, Corning, NY). ART 10 and 20P pipette tips (Molecular BioProducts, San Diego, CA) (see Note 2). 2. 200 mM bis-tris propane (Sigma, St. Louis, MO). 3. 15 mM triscarboxyethylphosphine (TCEP; Sigma, St. Louis, MO), pH 7.5. 4. 100 mg/mL fatty acid-free BSA. 5. 10X inhibitor stock (see Note 4). 6. 10 μM LpxC, iolated as described (6,13) (see Note 5). 7. 13 μM [14 C]-UDP-3-O-(R-hydroxymyristoyl)-N-acetyl-glucosamine. Store all buffers and reagents at –20 °C, and store LpxC at –80 °C. 8. 1.25 M NaOH. 9. 80:20 iso-propanol (iPrOH)/1.5 M acetic acid. 10. 0.1 M guanidine hydrochloride. 11. Scintiverse scintillation fluid (Fisher Scientific). Store at room temperature. Aliquot quench (1.25 M NaOH) into microfuge tubes prior to starting assay (see Note 6). 12. PEI-cellulose TLC plates (20 cm × 20 cm; Fisher Scientific) cut into 10 cm × 20 cm plates prior to use. 13. Biomax MS film, Kodak Photo Developer, and Kodak Photo Fixer. Dilute photo developer and fixer solutions with water (as indicated) prior to use.

2.3. Ultrafiltration Binding Assay 1. 200 mM bis-tris propane, 15 mM TCEP, pH 7.5 (10X buffer). Store at 4 °C (short term) or –20 °C (long term). 2. 0.1 M NaOH and Scintiverse scintillation fluid. Store at room temperature. 3. Microcons (MWCO 30K, Millipore, Bedford, MA). Microcons must be washed prior to assay to remove glycerol from the membranes (see Note 7) . 4. LpxC.

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5. Inhibitor. 6. [14 C]-UDP-3-O-(R-hydroxymyristoyl)-N-acetyl-glucosamine (see Notes 4, 5, 8). 7. Costar 3620 1.7 mL microcentrifuge tubes, Costar 4826 (0.1–10 μL) tips, and Costar 4863 (1–200 μL) tips (Corning Incorporated, Corning, NY). ART 200 and 1000 tips (Molecular BioProducts, San Diego, CA) (see Note 2).

3. Methods 3.1. Preparation of [14 C]-UDP-3-O-(R-hydroxymyristoyl) -N-acetyl-glucosamine 1. Mix together 100 μL 0.4 M HEPES, pH 8, 100 μL 10 mg/mL BSA, 540 μL H2 O, 200 μL [14 C]-UDP-N-acetyl-glucosamine, 10 μL 100 mM MMTS, and 30 μL 2–3 mM R,S-3-hydroxymyristate-ACP in 1.7 mL microfuge tube. Spin in microfuge for 10 s (see Note 9). 2. Initiate reaction with the addition of 20 μL 11–39 mg/mL LpxA to assay mixture. Vortex. Spin in microfuge for 10 s. Incubate in heat block at 30 °C for 30 min. 3. Transfer 1 μL of assay mixture into a scintillation vial (for counting). Load remaining assay mixture onto pre-equilibrated DEAE column. 4. Wash column with 18 mL 0.2 M NaCl to load radioactivity onto sep-pak. Remove sep-pak from DEAE-sepharose column. Discard DEAE column in radioactive waste. 5. Wash sep-pak with 30 mL H2 O using a disposable luer-lok syringe to remove unreacted [14 C]-UDP-N-acetylglucosamine. Collect waste in a 50 mL conical tube, and dispose of in radioactive waste (if necessary). 6. Elute [14 C]-UDP-3-O-(R,S-hydroxymyristoyl)-N-acetyl-glucosamine into 50 mL conical tube with 15 mL methanol using a disposable luer-lok syringe. 7. Concentrate methanol solution to dryness using a speed vac (see Note 9). 8. Resuspend [14 C]-UDP-3-O-(R-hydroxymyristoyl)-N-acetyl-glucosamine in 100 μL 10 mM bis-tris pH 6, and transfer into 1.7 mL microfuge tube. Transfer 1 μL of product into scintillation vial (for counting). Store [14 C]-UDP-3-O-(Rhydroxymyristoyl)-N-acetyl-glucosamine at –20 °C. 9. Add scintillation fluid to vials and count using scintillation counter. Determine reaction yield from the ratio of purified product (cpm)/total assay mixture (cpm). Yields for purified [14 C]-UDP-3-O-(R-hydroxymyristoyl)-N-acetyl-glucosamine using this procedure are typically >90%. Concentrations of stored product can be determined using the specific activity of purchased [14 C]-UDP-N-acetyl-glucosamine.

3.2. LpxC Deacetylase Turnover Assay for Measuring Inhibition Constants 1. Mix together 20 mM bis-tris propane, 1.5 mM TCEP pH 7.5, 1 mg/mL BSA with or without inhibitor in 1.7 mL microfuge tube. Spin in microfuge. Equilibrate assay

A Method to Assay Inhibitors of Lipopolysaccharide Synthesis

2.

3.

4.

5.

6. 7.

8.

9. 10. 11. 12. 13. 14. 15.

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mixtures at desired temperatures, 30 °C for E. coli LpxC (13) or Pseudomonas aeroginosa LpxC (8) or 60 °C for Aquifex aeolicus LpxC (17,18), for at least 15 min prior to initiating reactions. In a separate 1.7 mL microfuge tube, dilute 10 μM LpxC to 30 nM, so that the final solution contains 20 mM bis-tris propane, 1.5 mM TCEP pH 7.5, and 1 mg/mL BSA. Add diluted LpxC to assay mixture (with or without the inhibitor) pre-equilibrated to the assay temperature (30 or 60 °C). Vortex. Incubate this solution at the assay temperature for 15 to 25 min as needed to form the E•I complex. Initiate reaction with the addition of 13 μM [14 C]-UDP-3-O-(R-hydroxymyristoyl)N-acetyl-glucosamine to assay mixture at a final concentration of 200 nM. Vortex. For determination of kcat -, K M -, and KI -values, the initial linear rate needs to be determined at multiple concentrations of substrate (i.e., 50 nM to 8 μM) and/or inhibitor. Remove aliquots after various times (≥ 6 different time points) and add to quench (see Note 6). For reactions without inhibitor, recommended time points are every 30 s, whereas reactions that are significantly slowed by the addition of inhibitor will require longer time points (i.e., every 1–5 min) for better accuracy. Let quenched aliquots sit at room temperature for at least 10 min. Neutralize quenched aliquots with the addition of 80:20 iPrOH/1.5 M acetic acid and vortex (see Note 6). Concentrate neutralized aliquots down to ∼2 μL using speed vac. Resuspend concentrated aliquots in 14 μL 10 mM bis-tris, pH 6. Spin in microfuge for 30 s. Spot mixture onto PEI-cellulose plates in two portions. Let plates sit at room temperature until dry (∼30 min). Soak plates in methanol for 5 min each. Let sit at room temperature until dry (∼ 5 min). Run plates in 0.1 M guanidine hydrochloride until solvent front reaches the top of plate (∼11 min). Let plates sit at room temperature until dry (∼1 h). Expose plates to film, and place in film cassettes. Place at –80 °C overnight. Develop film. Cut substrate and product from TLC plates, and transfer into scintillation vials (see Note 10). Add scintillation fluid. Count substrate and product using scintillation counter. Convert fraction product (cpmproduct /cpmtotal ) into [product]. Plot [product] as a function of time, and fit linear equation to data to obtain initial linear rate (v; Fig. 1). Plot observed velocity (v) as a function of inhibitor concentration ([I]) and fit Eq. (1) to these data to obtain IC50 values (Fig. 2). For determination of kcat -, KM -, and KI -values, plot observed activity (v) as a function of substrate concentration ([S]) and fit Eq. (2) to these data:

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Fig. 1. Example of initial linear product formation for turnover of [14 C]-UDP-3-O(R,S-hydroxymyristoyl)-N-acetyl-glucosamine catalyzed by EcLpxC (20 mM bis-tris propane, 10 mM TCEP, pH 7.5, 30 °C).

Fig. 2. Example of LpxC inhibition by a hydroxamate inhibitor (20 mM bis-tris propane, 10 mM TCEP, pH 7.5, 30 °C). Initial linear rates of LpxC turnover were plotted as a function of concentration inhibitor, and Eq. (1) was fit to these data to obtain an IC50 of 3.6 ± 0.4 nM.

A Method to Assay Inhibitors of Lipopolysaccharide Synthesis v=

IC50  I + IC50 

v= KM



Vmax S   1 + KI + S

149 (1)

(2)

I

3.3. Ultrafiltration Binding Assay 1. For direct binding measurements, mix together 10X buffer, H2 O, LpxC (varied concentrations, i.e., 1–200 μM), and inhibitor (constant concentration, i.e., [I]/KI = 0.1) in 1.7 mL microfuge tubes so that the final concentration of buffer is 1X. For competition-based assays, [14 C]-UDP-3-O-(R-hydroxymyristoyl)-N-acetylglucosamine (73 nM) is also included in reaction mixtures. Spin in microfuge (see Notes 8 and 11). 2. Incubate at 30 °C for 15 to 30 min to allow for ligand equilibration/product formation. 3. Transfer mixtures into pre-rinsed microcons (see Note 7). 4. Spin at 3000X g for 3 min. Transfer filtrate back into sample reservoirs. Vortex. This step is to ensure that the filtrate sample is not diluted by residual buffer from the washing steps. 5. Spin at 3000X g for 2.5 min. Transfer 75 μL each of filtrate into scintillation vials. 6. Place sample reservoirs containing the retentate upside down in new collection tubes. Spin at 3000X g for 3 min. Transfer 75 μL each into scintillation vials. 7. Add scintillation fluid to vials and count using scintillation counter. 8. Plot EL/Ltotal , where L = radiolabeled ligand, as a function of [LpxC] (Fig. 3). KD Inhibitor -values are obtained by fitting Eq. (3) to these data for direct binding assay, or Eq. (4) for competition-based assays, where KD Product refers to the KD value for [14 C]-UDP-3-O-(R-hydroxymyristoyl)-glucosamine binding to EcLpxC under similar conditions (i.e., 1.5 ± 0.2 μM; Fig. 3).

 EL  Ltotal Endpt   EL  + EL L =   total Background Ltotal 1 + EKD

(3)

   EL    Ltotal  EL  Pr oduct    + EL L Initial  = KD Ltotal 1 + LpxC 1 + Inhibitor K Inhibitor

(4)



total

D

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Fig. 3. Example of [14 C]-UDP-3-O-(R,S-hydroxymyristoyl)-glucosamine binding to EcLpxC (20 mM bis-tris propane, 1.5 mM TCEP, pH 7.5, 30 °C) determined using ultrafiltration. Equation (3) was fit to these data to obtain a KD Product -value of 1.5 ± 0.2μM.

4. Notes 1. All water used in the described methods is milliQ purified water. 2. LpxC is catalytically active with one bound Zn(II) (E•Zn); however, LpxC can bind a second Zn(II) to form an inactive E•Zn2 complex (6,17). Therefore, lowmetal-content tubes and tips are used throughout the methods to minimize Zn(II) contamination and formation of E•Zn2 during assays and storage. Aerosol-resistant tips are used for pipetting radioactive materials. 3. Procedures for preparation of R,S-3-hydroxy-myristoyl-ACP and LpxA are provided in references (24,25). 4. Inhibitor stocks should be prepared at 10X the desired final concentration. Watersoluble LpxC inhibitors should be dissolved in 1X buffer. DMSO (≤ 10 % v/v final) can be included for inhibitors with low water solubility. 5. To minimize formation of E•Zn2 , all metal ions should be removed from purified LpxC by treatment with 20 mM dipicolinate as previously described (6). The catalytic Zn(II) is added back by incubation of apo-LpxC with Zn(II) (6). The concentration of LpxC required will depend on the type of assay, as well as the inhibitor potency. For assays that measure deacetylase activity, low concentrations of LpxC are typically required (i.e., 1–10 nM), while much higher concentrations of LpxC are required for assays that measure binding affinity (i.e., 1–200 μM).

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6. The amount of 1.25 M NaOH required will depend on the volume of the reaction being quenched. For a typical assay, 20 μL of reaction is quenched with 8 μL 1.25 M NaOH prior to neutralization with 94 μL 80:20 i-PrOH/1.5 M acetic acid. 7. To wash microcons, fill sample reservoirs with 500 μL each of 0.1 M NaOH. Spin at 14,000X g for 12 min. Discard filtrate. Repeat for a total of three washes. Fill sample reservoirs with 500 μL 25 mM bis-tris propane, 1.5 mM TCEP, pH 7.5. Spin at 14,000X g for 12 min. Discard filtrate. Fill sample reservoirs with 500 μL 20 mM bis-tris propane, 1.5 mM TCEP, pH 7.5. Microcons can be stored in buffer at 4 °C for several days prior to use. Immediately before use, spin at 14,000X g for 12 min. Place the sample reservoirs upside down over collection tubes. Spin at 3000X g for 3 min. Discard filtrate. Insert sample reservoirs into collection tube. Microcons are now ready for loading samples. 8. The KD -values of radiolabeled, or fluorescent, inhibitors for LpxC are determined using a direct binding assay, while the KD -values of unlabeled inhibitors are determined using a competition-based assay between the unlabeled inhibitor and [14 C]-UDP-3-O-(R,S-hydroxymyristoyl)-glucosamine. In these experiments, the concentration of either LpxC or inhibitor is varied, and the concentration of the other reagent held constant (below the KD -value). Since LpxC overexpresses at high levels and is unstable at low concentrations, the concentration of LpxC is typically varied in these experiments (typical range 1–200 μM). Both direct and competition-based assays require EL/Ltotal measurements at 6 to 10 different concentrations of LpxC to obtain a KD -value for a specific ligand. 9. The 3-acyl group on UDP-3-O-(R-hydroxymyristoyl)-N-acetyl-glucosamine can undergo isomerization to afford a molecule that is not a substrate for LpxC (24). To avoid isomerization of the acyl group, it is important to carry out the synthesis as quickly as possible, and the reaction should be kept cool. Therefore, the DEAEsepharose columns and sep-paks should be equilibrated prior to initiating the LpxA reaction, a syringe should be used to speed the rate of flow through the column, and the speed vac concentration step should be carried out at room temperature. If materials needed for the biochemical preparation are not readily available, radiolabeled UDP-3-O-(R-hydroxymyristoyl)-N-acetyl-glucosamine can also be prepared synthetically (26). 10. The length of film exposure will depend on the amount of radioactivity loaded onto the TLC plates, but in general ≥16 h is long enough to visualize substrate and/or product. Substrate migrates ∼1 cm below product on the PEI-cellulose TLC plate (Rf -values of ∼0.5 and ∼0.6, respectively), under these conditions. In experiments where product formation is low (steady-state conditions, 125 μM by broth MIC were next tested for synergy with azithromycin and linezolid by one concentration synergy tests.

3.4. MIC Synergy Testing in the Presence of EPI In strains containing efflux pumps, a successful EPI, in combination with azithromycin or linezolid, will allow the antibiotic in an in vitro susceptibility test to have a lower MIC than in the absence of the inhibitor. This test measures the change in MIC due to the presence of a constant concentration of EPI. Strains used for this test were EC1639, EC1764, and 54A1116, all strains containing the acrAB efflux pump. One 96-well U-bottom plate was used for two efflux pump inhibitors at 50 μM in combination with azithromycin and linezolid in a range of two-fold concentrations above and below the MICs determined in the previous study. 1. As described above, strains were grown from frozen stocks on the appropriate agar plates, for 2 nights before use, with passage of each strain onto a fresh agar plate after one night. 2. Using the appropriate media for the organism, each 96-well plate was filled with 190 μL of media in wells A1–H1. The remaining wells were filled with 95 μL of media. 3. Antibiotic at a concentration that was 20X the final concentration needed in the first well was dissolved in DMSO. The final concentration of antibiotic should be 1 to 2 dilutions above the actual MIC for that compound with that organism. 4. Ten μL of the 20X azithromycin were added to wells A1 and E1. Ten μL of the 20X linezolid were added to wells B1 and F1. Ten μL of DMSO were added to wells C1, D1, G1, and H1. 5. 1:2 serial dilutions of 100 μL from column 1 to column 11 were performed by a Biomek 2000, removing the last 100 μL from the plate. Column 12 received only 95 μL of media and served as the growth control wells. Plates now contained 95 μL of antibiotic at 1X the final concentration in two-fold serial dilutions. 6. The density of a bacterial culture was adjusted to an OD625 of 0.1, as read on a spectrophotometer, and then diluted 1:10. A multichannel pipette was used to

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inoculate the plates with 5 μL of this diluted culture for a final organism concentration of approximately 5 × 105 CFU/mL. One organism inoculated rows A–D, while a second organism inoculated rows E–H. 7. Added to the inoculum was the test EPI at a concentration of 10 mM. 38 μL of EPI at 10 mM were added to 340 μL of 1:10 diluted inoculum, and then 5 μL of inoculum were added to the 95 μL in the plate, for a 200-fold dilution of EPI. The final concentration of EPI in the well was 50 μM. 8. The plates were incubated at 35°C, ambient air, for 18 ± 2 h. 9. MICs of antibiotic in the presence of 50 μM of EPI were read by eye. Any compound that had an eight-fold or larger decrease in the MIC of the antibiotic in the presence of inhibitor compared to the MIC of the antibiotic alone, in at least one of the three strains tested, was further tested by FIC testing.

3.5. FIC Testing The most frequently used method to test antibiotic combinations in vitro is the fractional inhibitory concentration (FIC) method, also known as the checkerboard method. The checkerboard method is so named because of the pattern of wells formed by multiple dilutions of two compounds being tested, at concentrations above, at, and below their MICs (12). The results of a checkerboard test after 22 h of incubation provide a static view of the inhibition of growth of the organism in the presence of antibiotic in combination with the EPI. These results can demonstrate synergy, indifference, or antagonism between the two compounds. Each potential EPI was tested in combination with azithromycin or linezolid at concentrations starting at two or three dilutions above each compound’s MIC in a 96-well plate. The FIC for each drug is derived by dividing the lowest concentration of that drug in combination that inhibits growth, the MIC, by the MIC of that drug alone. Calculations from the MICs of each combination in a row of the plate gave an FIC index. The FIC index is the sum of the two FICs for each drug in combination (12). Synergy is defined as an FIC index less than or equal to 0.5, indifference is defined as an FIC index between 0.5 and 4.0, and antagonism is defined as an FIC index greater than 4.0. Strains used for FIC testing were EC1639, EC1764, and 54A1116. 1. As described above, strains were grown from frozen stocks on the appropriate agar plates, for two nights before use, with passage of each strain onto a fresh agar plate after one night. 2. One 96-well U-bottom plate was used for each efflux pump inhibitor/antibiotic combination. Each 96-well plate was filled with 25 μL of media, appropriate for the organism, in wells A1–H8. This 8 × 8 grid will contain the two compounds in combination.

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3. A11 through H11 and A12 through H12 were filled with 50 μL of media. These wells will determine the MIC of the inhibitor and the antibiotic alone, on the same plate as the compounds in combination. 4. Test compounds were dissolved at 40 mg/mL in DMSO. 5. Compound stocks were further diluted in the appropriate media to 4X and 8X the final maximum concentration in the plate. 6. 50 μL of the EPI, at 4X the final concentration, were added to well A11, and 50 μL of the antibiotic, at 4X the final concentration, were added to well A12. 7. Wells A11 and A12 were serially diluted 1:2 through wells H11 and H12 by a Denley Wellpro (Progroup Inc, St. Louis, MO), with the remaining 50 μL removed from the plate. 8. 25 μL of the EPI at 8X the starting concentration was added to wells A1 through H1. The drug was serially diluted 1:2 through column 8, with the remaining 25 μL removed from the plate. 9. The antibiotic was made at 4X the starting concentration in 10 mL of media. 10. Seven 1:2 serial dilutions in 5 mL were made. 11. Using an 8-channel pipette, 25 μL of the highest concentration of antibiotic were added to wells A1–A8. 12. The next-highest concentration was added to wells B1–B8, etc. The final volume of drug was 50 μL in each well, at a concentration of 2X the final concentration (Table 1). 13. To inoculate a plate, several colonies of each strain were added to a 5-mL flatbottom saline tube and mixed for 5 s, to an OD of 0.5 measured in a Crystalspec nephelometer. This gave a cell density of approximately 1 × 108 CFU/mL. 14. Each inoculum was diluted 1:100 in HTM or CAMHB. 15. 50 μL of diluted inoculum were added to each well, for a final organism density of 5 × 105 CFU/mL, in a final volume of 100 μL, and a drug concentration of 1X the final concentration. 16. The 96-well plates were incubated at 35°, ambient air, for 22 h and read by eye. 17. FIC indexes were determined by the following formula: (MIC of antibiotic in combination/MIC of antibiotic alone) + (MIC of EPI in combination/MIC of EPI alone). 18. An alternative method of making checkerboard plates is described in Section 3.8.

3.6. Protonophore Assay Any active EPI that demonstrated synergy by FIC was tested in the protonophore assay to rule out that its activity was due to disruption of the proton motive force (PMF). Protonophores disrupt cell membrane potentials and therefore dissipate the PMF across the membranes, depriving the efflux pumps of energy. The constructed E. coli strain JL467 will accumulate 14 Clactose within its cells, unless the PMF is disrupted by a test compound. CCCP,

Table 1 Example of Drug Concentrations (μg/mL) in a 96-Well Plate Alone and in Combination EPI/ Antibiotic A B C D E F G H

1

2

3

4

5

6

7

8

64/8 64/4 64/2 64/1 64/0.5 64/0.25 64/0.125 64/0.0625

32/8 32/4 32/2 32/1 32/0.5 32/0.25 32/0.125 32/0.0625

16/8 16/4 16/2 16/1 16/0.5 16/0.25 16/0.125 16/0.0625

8/8 8/4 8/2 8/1 8/0.5 8/0.25 8/0.125 8/0.0625

4/8 4/4 4/2 4/1 4/0.5 4/0.25 4/0.125 4/0.0625

2/8 2/4 2/2 2/1 2/0.5 2/0.25 2/0.125 2/0.0625

1/8 1/4 1/2 1/1 1/0.5 1/0.25 1/0.125 1/0.0625

0.5/8 0.5/4 0.5/2 0.5/1 0.5/0.5 0.5/0.25 0.5/0.125 0.5/0.0625

9 10 EPI Antibiotic — — — — — — — —

— — — — — — — —

64 32 16 8 4 2 1 0.5

8 4 2 1 0.5 0.25 0.125 0.0625

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a known proton motive force inhibitor, and cycloserine, which has no effect on membranes, are included in the assay to validate the assay conditions. LB broth serves as a positive control, allowing full uptake of the radiolabel. Any EPI tested at 3X the MIC that had less than 50% 14 C-lactose accumulation after 30 min, compared to LB broth alone, was eliminated. 1. A 10 μL loop JL467 frozen stock was grown in LB broth containing 100 μg/mL of ampicillin at 37°C, in ambient air, overnight. Overnight culture was diluted 1:100 in LB broth containing 100 μg/mL of ampicillin. This culture was grown for 3 h, until the OD600 read on a spectrophotometer equaled 1.0. 2. One 96-well flat-bottom plate can test 15 compounds at three concentrations, 1X, 2X, and 3X the MIC of that compound, in duplicate. Also included on the plate is CCCP, at 5 μg/mL final concentration, cycloserine, at 50 μg/mL final concentration, and two wells that contain the positive control, LB broth. 3. In Eppendorf tubes, 100 μL of each compound at 12X the final concentration were prepared in LB broth. Also, prepared were 100 μL of CCCP in LB broth at 60 μg/mL and 100 μL of cycloserine in LB broth at 600 μg/mL. 4. Compound was added to 96-well flat-bottom plates in duplicate by adding 25 μL of the highest concentration of compound 1 to wells A1 and B1, the second-highest concentration in wells A2 and B2, and the lowest concentration of compound in wells A3 and B3. The remaining wells were filled similarly with compounds 2 through 15. 25 μL CCCP were added to wells G10 and H10, 25 μL of cycloserine were added to wells G11 and H11, and 25 μL of LB broth were added to wells G12 and H12. 5. 25 μL of 14 C-lactose in LB broth were added to all wells. 6. Following the 3-h incubation of cells, to an OD600 = 1.0, 200μL cells were added to each well. 7. The plate was incubated at 37°C for 30 min. 8. The reaction was stopped by transferring the entire volume of each well to a GF/B filter plate and filtering the contents by a Filtermate harvester (PerkinElmer, Boston, MA). The plate was washed 3 times with 200 μL of cold stop solution. 9. The plate was air-dried overnight. 10. The bottom of plate was covered, and 25 μL of liquid scintillation cocktail were added to each well. The top of the plate was covered, and the plate was counted for 14 C in 1-min intervals in a Wallac Trilux liquid scintillation counter (EG&G Wallac, Gaithersburg, MD). 11. Use the counts from the wells containing only LB broth as 100% uptake, and normalize all other counts to percent uptake compared to LB broth. As an active protonophore, CCCP will have less than 50% uptake. Cycloserine will have >80% uptake. At a concentration that is 3X the MIC, a potential EPI that is not protonophore-like will have more than 60% uptake of 14 C-lactose.

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3.7. Kill Kinetic Studies Kill kinetic studies are used to demonstrate the rate of killing of an antibiotic over time. An effective inhibitor would increase the rate of killing if it inhibited the efflux pumps and allowed the antibiotic to stay in the cell. EPIs were tested at 50 and 25 μM along with an untreated control against azithromycin at concentrations above and below the MIC for azithromycin in that strain, over 24 h. Cell counts at each time point were plotted in log scale versus time to look for a synergistic killing effect of the EPI in combination with azithromycin, compared to the rate of killing of azithromycin alone. Synergy is defined as more than 2 logs greater killing after 24 h compared to the best compound alone (10). Strains used for kill kinetics were EC1639 and EC1764. 1. Strains were grown as previously described on Tryptic soy agar plates overnight. Colonies were scraped off plates, transferred to 5 mL of CAMHB in 15-mL tubes, and incubated at 35°C the second night. In the morning, strains were diluted to OD625 of 0.12, and then diluted 1:100 in CAMHB, for a starting cell concentration of 1 × 106 CFU/mL. 2. Stock concentration of azithromycin was made in CAMHB to 1 mg/mL. Azithromycin was further diluted in CAMHB to 8.0 μg/mL, 4X the final concentration needed in the plate. Four 4-fold dilutions of the 8.0 μg/mL stock were made in CAMHB, in a final volume of 1 mL. 3. Each EPI at 100 mM, and 50 mM, 2X the final concentration in the plate, were made in CAMHB, to a volume of 1 mL. 4. One 24-well plate was used for each inhibitor/azithromycin/strain combination. 250 μL of the highest concentration of azithromycin were added to wells A1, B1, and C1. Each of the four additional dilutions of azithromycin was added to all wells in columns 2–5, respectively. Column 6 received 250 μL of CAMHB. 5. All wells in row A received 250 μL of inhibitor at the highest concentration. All wells in row B received the lower concentration of inhibitor. Row C received 250 μL of CAMHB. Row D was not used. 6. 500 μL of the diluted bacteria were added to all wells, A1–C6. The final volume in each well was 1.0 mL, the final concentration of azithromycin in column 1 was 2.0 μg/mL, and the final concentration of inhibitor in row 1 was 50 mM. 7. 24-well plates were incubated at 37°C, ambient air, for 24 h. At 0, 2, 4, 6, and 24 h, samples from each well were taken to determine colony counts. A 20-μL sample from each well was added to the first row of a 96-well plate previously filled with 45 μL of PBS in all wells except row 1. Row 1 was reserved for undiluted sample. Five μL of the undiluted sample were serially diluted to row 8. Two 96-well plates were needed to dilute the 18 different inocula in each 24-well plate. Using an 8-channel multichannel pipette, 10 μL from wells 1–8 from each column were pipetted onto an LB agar plate, in duplicate on the same plate, across a range of 102 to 109

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CFU/mL. Plates were allowed to sit for 10 min on the bench before incubating at 37°C, ambient air, overnight. Colonies at each dilution were counted and plotted on a log scale versus incubation time for each antibiotic/inhibitor combination.

3.8. Alternative Method of FIC Testing Using Alamar Blue For each EPI-antibiotic combination tested, rows A2 through H8 of one 96-well deep-well plate were filled with 400 μL of CAMHB broth containing 1% Alamar Blue by the Multidrop instrument (Labsystems, Helsinki, Finland). 1. Efflux pump inhibitors were dissolved in DMSO at 2.56 mg/mL. Compounds were further diluted to 16X the final concentration needed in CAMHB or HTM. Each potential efflux pump inhibitor was tested in combination with azithromycin or linezolid at concentrations starting at two or three dilutions above each compound’s MIC. 2. 800 μL of the EPI were added to wells A1–H1. Two-fold serial dilutions from column 1 to column 7 of each deep-well mother plate were performed by the Biomek 2000 instrument (Beckman-Coulter). The remaining 400 μL were removed from the plate. Column 8 contained no EPI drug. 3. The antibiotic was made at 16X the starting concentration in 10 mL of media. 4. Six 1:2 serial dilutions of the antibiotic in 5 mL were made. 5. 400 μL of the highest concentration of the antibiotic were then added to wells A1– A8 of each deep-well plate containing a specific EPI. 400 μL of the next-highest concentration of antibiotic were added to wells B1–B8, etc., through row G. Well H8 served as the growth control well. 6. There is now 800 μL total volume in each well. Wells A1 through G8 contain the EPI-antibiotic combination at 8X the final concentrations. Wells H1 through H7 contain the EPI only at 8X the final concentrations, and wells A1 through A8 contain the antibiotic only at 8X the final concentration (Table 2). 7. 25 μL were transferred by the Multimek 96 instrument (Beckman-Coulter) from each well of each mother plate to a daughter plate containing 165 μL of medium. Twelve daughter plates could be created from each mother plate. This was an eight-fold dilution of compound after the inoculum was added. 8. Strains were grown as described above. The Biomek instrument was used to inoculate the daughter plates with 10 μL of a 1:10-diluted culture equal to an OD625 of 0.1 as read on a spectrophotometer (approximately 1 × 108 CFU/mL), for a final organism concentration of approximately 5 × 105 CFU/mL. The plates were incubated at 35°C, ambient air, for 18 ± 2 h. 9. The plates were read visually by observing a reduction (a color change from blue to purple or pink, indicating organism growth) or no reduction [a blue color, indicating no growth of organism due to inhibition by a drug(s)]. FICs were determined by the following formula: (MIC of antibiotic in combination/MIC of antibiotic alone) + (MIC of EPI in combination/MIC of EPI alone).

Table 2 Example of Drug Concentrations (μg/mL) in a 96-Well Plate Alone and in Combination, Alternative Method EPI/Antibiotic A B C D E F G H

1

2

3

4

5

6

7

8

9

10

11

12

64/8 64/4 64/2 64/1 64/0.5 64/0.25 64/0.125 64

32/8 32/4 32/2 32/1 32/0.5 32/0.25 32/0.125 32

16/8 16/4 16/2 16/1 16/0.5 16/0.25 16/0.125 16

8/8 8/4 8/2 8/1 8/0.5 8/0.25 8/0.125 8

4/8 4/4 4/2 4/1 4/0.5 4/0.25 4/0.125 4

2/8 2/4 2/2 2/1 2/0.5 2/0.25 2/0.125 2

1/8 1/4 1/2 1/1 1/0.5 1/0.25 1/0.125 1

8 4 2 1 0.5 0.25 0.125 Growth

— — — — — — — —

— — — — — — — —

— — — — — — — —

— — — — — — — —

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4. Notes 1. Because E. coli has multiple antibacterial efflux pumps, two strains, EC1763 and EC1764, derived from parent strain EC1639, were constructed by Keith Poole (Queens University, Kingston, Ontario) to examine the effect of inhibiting only the acrAB pump. E. coli strain EC1763 had four chromosomally encoded efflux pumps knocked out as well as waaP (quad strain—acrAB, acrEF, emrE, emrD—with a waaP deletion). waaP is a kinase in the outer membrane of E. coli that is responsible for adding the LPS component to the outer membrane. Deletion of this kinase increases the outer membrane permeability of this strain, allowing potential efflux pump inhibitor compounds to more readily reach the efflux pumps. EC1763 was transformed with a plasmid vector encoding ampicillin resistance and the structural genes for acrAB and the new strain containing only acrAB was designated EC1764. EC1764 was grown on LB agar containing 100 μg/mL of ampicillin. The other related E. coli strains, EC1639 and EC1763, were grown on LB agar or Tryptic soy agar without added antibiotic. 2. Previous tests found the largest data variation occurred from inocula made on separate days. Therefore, replicates were run over three days instead of three plates on one day or three wells in one plate on a single day.

Acknowledgments We acknowledge the work of numerous former colleagues at Pharmacia who started this research program: D. E. Decker, M. P. Sheets, S. E. Buxser, M. R. Barbachyn, R. Gadwood, G. Zurenko, J. Bourdage, G. F. Hess, J. K. Gibson, A. G. Morgan, J. E. Mott, et al. We also thank Keith Poole and colleagues at Queens University in Kingston, Ontario, for E. coli strain constructs. References 1 Li, X.-Z., and Nikaido, H. (2004) Efflux-mediated drug resistance in bacteria. 1. Drugs 64, 159–204. 2 Yu, E. W., McDermott, G., Zgurskaya, H., Nikaido, H., and Koshland, D. E. Jr. 2. (2003) Structural basis of multiple drug-binding capacity of the AcrB multidrug efflux pump. Science 300, 976–980. 3 Yu, E. W., Aires, J. R., and Nikaido, H. (2003) AcrB multidrug efflux pump 3. of Escherichia coli: Composite substrate-binding cavity of exceptional flexibility generates its extremely wide substrate specificity. J. Bacteriol. 185, 5657–5664. 4 Yu, E. W., Aires, J. R., McDermott, G., and Nikaido, H. (2005) A periplasmic 4. drug-binding site of the AcrB multidrug efflux pump: A crystallographic and site-directed mutagenesis study. J. Bacteriol. 187, 6804–6815. 5 Murakami, S., Nakashima, R., Yamashita, E., and Yamaguchi, A. (2002) Crystal 5. structure of bacterial multidrug efflux transporter AcrB. Nature 419, 587–593.

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6 Murakami, S., Nakashima, R., Yamashita, E., Matsumoto, T., and Yamaguchi, A. 6. (2006) Crystal structures of a multidrug transporter reveal a functionally rotating mechanism. Nature 443, 173–179. 7 Higgins, M. K., Bokma, E., Koronakis, E., Hughes, C., and Koronakis, V. (2004) 7. Structure of the periplasmic component of a bacterial drug efflux pump. Proc. Natl. Acad. Sci. USA 101, 9994–9999. 8 Anonymous (2003) National nosocomial infections surveillance (NNIS) system 8. report, data summary from January 1992 through June 2003, issued August 2003. Amer. J. Infect. Control 31, 481–498. 9 Baker, C. N., Banerjee, S. N., and Tenover, F. C. (1994) Evaluation of alamar 9. colorimetric MIC method for antimicrobial susceptibility testing of Gram-negative bacteria. J. Clin. Microbiol. 32, 1261–1267. 10 Clinical and Laboratory Standards Institute. (2006) Methods for Dilution Antimi10. crobial Susceptibility Tests for Bacteria That Grow Aerobically; Approved Standard – Seventh Edition. CLSI Document M7-A7. 11 Amsterdam, D. (1996) Susceptibility testing of antimicrobials in liquid media. In 11. Antibiotics in Laboratory Medicine, 4th ed. (V. Lorian, ed.). The Williams and Wilkins Co., Baltimore, MD, pp. 52–111. 12 Eliopoulos, G. M., and Moellering, R. C. Jr. (1996) Antimicrobial combination. 12. In Antibiotics in Laboratory Medicine, 4th ed. (V. Lorian, ed.). The Williams and Wilkins Co., Baltimore, MD, pp. 330–396.

16 Mycobacterium tuberculosis -Ketoacyl Acyl Carrier Protein Synthase III (mtFabH) Assay: Principles and Method Sarbjot Sachdeva and Kevin A. Reynolds

Summary Fatty acid biosynthesis is one of the relatively newer targets in antibacterial drug discovery. The presence of distinct fatty acid synthases (FAS) in mammals and bacteria and the fact that most bacterial FAS enzymes are essential for viability make this a very attractive antimicrobial drug target. The enzyme -ketoacyl ACP synthase (KASIII or FabH) is the key enzyme that initiates fatty acid biosynthesis in a type II dissociated FAS. This enzyme catalyzes the condensation of acyl CoA and malonyl ACP (acyl carrier protein) to form a -ketoacyl ACP product, which is further processed to form mature fatty acids that are involved in various essential cellular processes and structures like phospholipid biosynthesis, cell wall formation, etc. Herein we describe a new assay for the Mycobacterium tuberculosis FabH (mtFabH) enzyme involved in a key initiation step in the synthesis of mycolic acids, which are an integral component of the cell wall. The assay eliminates the need for the cumbersome washing steps or specialty scintillation proximity assay beads and the preparation of acyl carrier proteins required in other assay formats. This discontinuous assay involves the reduction of radiolabled long-chain -ketoacyl CoA product to its dihydroxy derivative, which partitions into a nonpolar phase for quantitation, while the reduced radiolabeled substrate derivative remains in the aqueous phase.

Key Words: FAS; fatty acid biosynthesis; lipid metabolism; mtFabH; -ketoacyl ACP synthase; KASIII; fatty acid condensing enzymes.

1. Introduction Bacterial fatty acid biosynthesis is an essential cellular process and has recently gained interest as a relatively unexplored drug target. In mammals, this From: Methods in Molecular Medicine, Vol. 142: New Antibiotic Targets Edited by: W. Scott Champney © Humana Press Inc., Totowa, NJ

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process is carried out by a single large polypeptide (a type I FAS) containing catalytic domains for all of the reactions and an integral acyl carrier protein (ACP) that ferry the substrates and products between them. Conversely, bacteria and plants use a type II FAS in which each reaction is carried out by separate enzymes. This distinct difference in mammalian and bacterial FAS systems suggests the possibility of agents that selectively inhibit only one process. Indeed, a number of antibacterial agents that target the type II FAS have been discovered (for comprehensive reviews, see Refs, 1–3). The enzymes involved in bacterial and plant type II FAS system and their known inhibitors are shown in Fig. 1. FabH catalyzes the first condensation reaction between an acyl CoA and malonyl ACP (MACP). Typically, this enzyme uses acetyl CoA or other short-chain acyl CoA substrates to initiate the biosynthesis of palmitoyl CoA and related fatty acids. In Mycobacterium tuberculosis, these long-chain fatty acids are generated by a type I FAS. A type II FAS elongates them further to form very long fatty acid chains (>C54 ), which are

Fig. 1. Roles of individual enzymes in a type II fatty acid synthase. Enzymes inhibited by triclosan, cerulenin, isoniazid, and TLM (thiolactomycin) are indicated (wavy lines).

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then elaborated to mycolic acids, a key structural component and permeability barrier of the M. tuberculosis cell wall. Accordingly, the M. tuberculosis FabH (mtFabH) has a different and unusual specificity for longer acyl CoA substrates. Convenient assays for this enzyme permit discovery and development of selective inhibitors and a potential new treatment for tuberculosis. 1.1. Assay Overview and Principles The most commonly used FabH assay involves the use of radioactive acyl CoA and MACP to form a radioactive -ketoacyl-ACP product. This radioactive protein (ACP) product can be precipitated with acid, washed, and counted, while the radioactive substrate stays in aqueous phase. The accuracy of the assay requires extensive time-consuming washing steps. Nonetheless, this general approach has been used to assay FabH from various pathogenic microorganisms such as E. coli, E. faecalis, H. influenzae, S. pyogenes, and S. aureus (4,5). Unfortunately, this method cannot be used to assay mtFabH, because the long-chain acyl CoA substrates utilized by mtFabH have poor solubility and are not readily resolved from the  -ketoacyl–ACP product by the acid precipitation and extensive washing steps. Other assay formats such as a scintillation proximity assay (SPA) (6) and a filter disk (7) assay have been described and can successfully be used for mtFabH. However, these assays involve either tedious preparation of a biotinylated malonyl ACP substrate (SPA) or numerous washing steps (filter disk assay). Recently, Brown et al. (8) developed an alternative method similar to an assay used originally by Garwin et al. (9) for FabF/FabB enzymes. This assay (Fig. 2) involves the reduction of the assay mixture after reaction completion to produce radioactive substrate and product derivatives that differ dramatically from each other in terms of polarity. This difference permits a simple separation of the two and quantitation of the radioactive product. The assay uses FabD to produce radioactive 2-14 C MACP in situ from holo-ACP and malonyl CoA. The mtFabH catalyzes a decarboxylative condensation of this MACP with the long-chain acyl CoA substrate (specificity ranging from hexanoyl CoA to arachidonoyl CoA; see Table 1) to form a long-chain  -ketoacyl ACP. Reduction of this product by NaBH4 gives the corresponding radioactive long-chain alkyl-1, 3-diol product with a 14 C label at C2. This product derivative can be partitioned into non-aqueous phase, whereas the radioactive propane-1,3-diol obtained by reduction of the radioactive malonyl thioesters stays in an aqueous phase. As described below, this assay can be simplified by using malonyl CoA directly as a substrate, instead of MACP (see Fig. 2).

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Fig. 2. Schematic of mtFabH assay. The principle of separation of radioactive substrate and product is described in Section 1.2. The radioactive center is shown by the asterisk *. R’ can be either CoA or ACP.

A slower reaction rate for mtFabH is observed when malonyl CoA is used in place of a MACP substrate (8) (see Table 1). Dodecanoyl CoA is the preferred substrate for assays using either malonyl CoA or ecMACP (generated from the Escherichia coli ACP). With this acyl CoA substrate, the reaction rate is about four-fold slower with malonyl CoA than with ecMACP or mtMACP (generated from the M. tuberculosis ACP). Catalysis of longer-chain acyl CoA substrates by the mtFabH is markedly reduced using either ecMACP or malonyl CoA, but not mtMACP. One advantage of this modified assay is that all the substrates are commercially available. There is no need to obtain a purified ACP, or to convert it to either the corresponding MACP or biotinylated MACP, leading to significant savings in both time and cost. The direct utilization of radioactive malonyl CoA in the assay also maximizes the use of this substrate for the assay, avoiding the inevitable losses associated with conversion to malonyl ACP. The assay also uses lower concentrations of malonyl CoA (12.5 μM vs. the 50 μM typically used in other formats (7,8), decreasing the amount of material and potentially increasing the range of the assay to include both poor

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Table 1 Specific Activity of mtFabH with Malonyl CoA (in Place of MACP) Using a Range of Different Acyl CoA Substrates Acyl CoA Primer Chain Length

Specific Activity mtFabH (nMoles/min/mg)

C6 C8 C10 C12 C14 C16 C18 C20

0.38 +/- 0.064 0.69 +/- 0.18 0.758 +/- 0.095 1.77 +/- 0.041 0.3528 +/- 0.0269 0.128 +/- 0.086 0.036 +/- 0.0009 0.155 +/- 0.0133

and good competitive inhibitors. A final advantage of this new assay format is an excellent 60:1 signal-to-noise ratio (compared with 10:1 in the scintillation proximity assay (6)). 2. Materials 1. Lauroyl CoA or other long-chain acyl CoA (Sigma)—Make a stock solution of 100 mM or 10 mM in aqueous hydrochloric acid, pH 3.5, then dilute to 100 μM working concentration and aliquot into appropriate working volumes (see Note 1). Store at –80 °C. 2. Radioactive [2-14 C] Malonyl CoA (1.8 mM; specific activity, 55 mCi/mmol, American Radiolabeled Chemicals, Inc.). Dilute approximately 7.2 times in aqueous hydrochloric acid solution (pH 3.5) and aliquot the resulting 250 μM solution into appropriate working volumes (see Note 2). Store at –80 °C. 3. mtFabH expression in E coli and purification has been described previously (6). 4. Sodium phosphate buffer: 100 mM sodium phosphate, pH 7.0, in double-distilled (DD) water or enzyme grade water. 5. Tetrahydrofuran (THF)—reagent grade. 6. Reducing reagent: 5 mg of NaBH4 to 1 mL of a 70% 100 mM K2 HPO4 , 100mM KCl, 30% THF solution. This solution is prepared fresh and kept on ice until used (10) (see Note 3). 7. Screw-cap microcentrifuge tubes. 8. Scintillation cocktail—EcoScint A (National Diagnostics). 9. Scintillation vials—Wheaton Science; high-density polyethylene (HDPE).

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3. Methods The reaction is carried out in 20 μL (total volume). This low volume using a minimum of the commercial radioactive malonyl CoA sample with an unchanged specific activity balances a high signal-to-noise ratio with a low cost. Larger-volume assays would require either additional radioactive malonyl CoA (increased cost) or the addition of unlabeled malonyl CoA (decreasing the specific activity and signal-to-noise ratio). The concentrations of various components in a typical assay are shown in Table 2. These volumes permit determination of an IC50 value for mtFabH inhibitor but can be modified to perform the other enzyme studies by replacement with an additional 2 μL of buffer. 1. Thaw the desired number of lauroyl CoA, malonyl CoA, and mtFabH aliquots and keep at 4 °C. 2. In a screw-cap microcentrifuge tube, add 2.5 μL of 100 μM lauroyl CoA solution (final concentration, 12.5 μM lauroyl CoA), 1 μL of the 250 μM [2-14 C] malonyl CoA solution (approx. 31,000 counts; final concentration, 12.5 μM), and 2 μL of 10X inhibitor concentration (for inhibitor IC50 studies). Add 0.1 M of sodium phosphate buffer, pH 7.0, to make up the volume to 18 μL. 3. Initiate the reaction by addition of 2 μL of mtFabH (0.2–0.4 μg) solution in 0.1 M of sodium phosphate buffer, pH 7.0. Cap the microcentrifuge tube and keep at 37 °C for a designated time (typically, 30–90 min). Under these assay conditions, a linear reaction rate was observed over a 2-h assay period (Fig. 3). 4. Add 0.5 mL of the reducing solution to the assay mixture, cap the vials, and shake vigorously. Incubate the mixture at 37 °C for 15 min (see Note 4). 5. Add 500 μL of toluene to the mixture, mix the solution vigorously, and allow to separate at room temperature (see Note 5). 6. Remove 320 μL of the upper phase (toluene), and combine with approximately 3 mL of scintillation cocktail. Table 2 List of Various Components of a Typical mtFabH Assay Component Lauroyl CoA Malonyl CoA Inhibitor (if any) mtFabH Sodium phosphate, pH 7.0 Total volume

Stock Conc.

Volume Used (μL)

Final Conc./Amount

100 μM 250 μM 10X 2 μg/μL 100 mM

2.5 1 2 2 12.5 20

12.5 μM 12.5 μM 1X 0.2 μg 72 mM

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10000

14C

Counts

8000

6000

4000

2000

0 0

20

40

60

80

100

120

Time in minutes

Fig. 3. Linearity of mtFabH activity with time. Enzyme is incubated with substrates at 37 °C for the specified time, and product formation is quantitated as described above. 7. Determine the radioactivity of the toluene extract with a scintillation counter, and double this value to obtain the actual radioactivity in non-aqueous phase (see Note 6).

4. Notes 1. As acyl CoA substrates are unstable at neutral and higher pH, the solutions are made in acidic pH (aqueous hydrochloric solutions at pH 3.5) and stored in aliquots at –80 °C in working volumes (avoiding the need to repeatedly thaw and refreeze the solutions and the associated increased degradation). 2. Malonyl CoA from this source comes in 1800-μM solution. It is diluted to a final concentration of 250 μM and stored at –80 °C at pH 3.5. The manufacturer’s technical data sheet states that under these conditions the rate of decomposition is approximately 2% over the first six months. A very important factor in this particular assay is the purity of the commercially available radioactive malonyl CoA. Radioactive malonyl CoA is prepared by the reaction of monothiophenylmalonate [malonyl-2-14 C] and coenzyme A, and the [2-14 C] malonyl CoA product is purified by column chromatography. The radioactive reactant is relatively nonpolar and, if present in the [2-14 C] malonyl CoA sample, can lead to a high background during the assay and a substantially reduced signal-to-noise ratio.

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3. Divalent cations like Mg2+ and Mn2+ catalyze the decomposition of sodium borohydride (NaBH4 ), preventing effective reduction of the 3-ketoacyl CoA product. Thus, the use of distilled water free of divalent ions is used in both the assay and the preparation of the reducing reagent solution (10). 4. Sodium borohydride produces significant effervescence when mixed with aqueous solutions. There is thus a potential for loss of the radioactive material from the vial at this point. A screw-cap microcentrifuge tube should be used and tightened carefully before shaking the sample. Reduction of long-chain acyl CoA is quantitative within 10 min. If inconsistent results are obtained, the reduction reaction can be carried out for a longer time. The THF in the reducing solution decreases the time required to quantitatively reduce the acyl thioesters (from 2 h to 10 min) (10). The buffer can be stored at room temperature, and NaBH4 should be added just before use. This reduction solution can be used for up to 2 h if kept on ice but potentially can lead to a loss of consistency with the assay results. 5. The volume of toluene solution increases to about 640 μL due to partitioning of THF into toluene. 6. Solvents used to deliver inhibitors have a predictable inhibitory effect on the activity on mtFabH. The effect of various concentrations of dimethylsulfoxide (DMSO) and methanol is shown in Fig. 4. This figure demonstrates that the concentrations of solvent should ideally be at or below 1% in the final assay volume. The order of substrate and inhibitor components can be changed to perform the assays with or without the enzyme-inhibitor preincubation. While this assay uses malonyl CoA (with the associated advantages discussed above), it can also be used with MACP.

120

DMSO MeOH

% mtFabH Activity

100 80 60 40 20 0 0.01

0.1

1

10

% Solvent

Fig. 4. Effect of solvent concentrations of dimethylsulfoxide (DMSO) and methanol (MeOH) on mtFabH activity.

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References 1 Heath, R. J., White, S. W., and Rock, C. O. (2001) Lipid biosynthesis as a target 1. for antibacterial agents. Prog. Lipid Res. 40, 467–497. 2 Rock, C. O., and Jackowski S. (2002) Forty years of bacterial fatty acid synthesis. 2. Biochem. Biophys. Res. Comm. 292(5), 1155–1166. 3 Heath, R. J., White, S. W., and Rock, C. O. (2002) Inhibitors of fatty acid synthesis 3. as antimicrobial chemotherapeutics. Appl. Microbiol. Biotechnol. 58, 695–703. 4 Nie, Z., Perretta, C., Lu, J., Su, Y., Margosiak, S., Gajiwala, K. S., Cortez, J., 4. Nikulin, V., Yager, K. M., Appelt, K., and Chu, S. (2005) Structure-based design, synthesis, and study of potent inhibitors of  -ketoacyl-acyl carrier protein synthase III as potential antimicrobial agents. J. Med. Chem. 48, 1596–1609. 5 Tsay, J. T., Oh, W., Larson, T. J., Jackowski, S., and Rock, C. O. (1992) Isolation 5. and characterization of the  -ketoacyl-acyl carrier protein synthase III gene (fabH) from Escherichia coli K-12. J. Biol. Chem. 267, 6807–6814. 6 Scarsdale, J. N., Kazanina, G., He, X., Reynolds, K. A., and Wright, H. T. (2001) 6. Crystal structure of the Mycobacterium tuberculosis -ketoacyl-acyl carrier protein synthase III. J. Biol. Chem. 276, 20516–20522. 7 Choi, K. H., Kremer, L., Besra, G. S., and Rock, C. O. (2000) Identification and 7. substrate specificity of -ketoacyl (acyl carrier protein) synthase III (mtFabH) from Mycobacterium tuberculosis. J. Biol. Chem. 275, 28201–28207. 8 Brown, A. K., Sridharan, S., Kremer, L., Lindenberg, S., Dover, L. G., 8. Sacchettini, J. C., and Besra, G. S. (2005) Probing the mechanism of the Mycobacterium tuberculosis -ketoacyl-Acyl carrier protein synthase III mtFabH; factors influencing catalysis and substrate specificity. J. Biol. Chem. 280(37), 32539–32547. 9 Garwin, J. L., Klages, A. L., and Cronan, J. E. Jr. (1980) Structural, enzymatic, and 9. genetic studies of -ketoacyl-acyl carrier protein synthases I and II of Escherichia coli. J. Biol. Chem. 255(24), 11949–11956. 10 Barron, E. J., and Mooney, L. A. (1968) Determination of acyl-thiolesters by 10. gas-liquid chromatography of their sodium borohydride reduction products. Anal. Chem. 40(11), 1742–1744.

17 Screening for Compounds That Affect the Interaction Between Bacterial Two-Component Signal Transduction Response Regulator Protein and Cognate Promoter DNA Matthew G. Erickson, Andrew T. Ulijasz, and Bernard Weisblum

Summary Bacterial signal transduction systems can be used as drug targets. The signal transduction targets fall into two groups—sensor kinases and response regulators. Previously reported studies describe hits that were thought to inactivate sensor kinases but on closer examination were found to act elsewhere instead; a possible reason for this is that full-length sensor kinases are integral membrane proteins whose activity might reflect interaction with the cell membrane or with membrane components. We describe a model system that instead is based on the interaction between a test compound and a response regulator in a homogeneous phase reaction. In this system, response regulator-DNA complex formation and its inhibition by a test compound are measured by fluorescence polarization. The model system should be readily adaptable to drug discovery based on other bacterial two-component s transduction systems.

Key Words: fluorescence polarization; signal transduction; sensor kinase; response regulator.

1. Introduction Two-component signal transduction (TCST) systems mediate adaptive changes in bacteria. The key participants are a sensor kinase and a response regulator. The response regulator protein is phosphorylated by its cognate activated sensor kinase and acts as transcription factor by binding to the promoter region of the gene(s) whose function it regulates. For reviews, see the studies by Hoch and Silhavy (1), Stock et al. (2), and West and Stock (3). Genetic From: Methods in Molecular Medicine, Vol. 142: New Antibiotic Targets Edited by: W. Scott Champney © Humana Press Inc., Totowa, NJ

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studies have shown that several TCST systems are essential for virulence or infectivity, suggesting that TCST systems may be suitable targets for screening compounds with anti-infective activity. Examples include (1) Streptococcus pneumoniae CiaRH (4) and RR489 (5), which control cephalosporin resistance and lung infectivity, respectively; (2) Mycobacterium tuberculosis MtrA, which is needed for infectivity (6); (3) Listeria monocytogenes CesRK, which regulates tolerance to ß-lactam antibiotics (7); and (4) Enterococcus faecium VanRS, which specifies inducible vancomycin resistance (8). Bacterial antibiotic resistance has become a problem that severely limits the clinical effectiveness of many of our commonly used antibiotics. One consequence has been the need for new antibiotics, which, in turn, has led to the search for new drug targets. One example of a group of potential drug targets is the TCST systems of bacteria. These systems generally consist of a sensor kinase and its cognate response regulator in which the kinase is associated with the cell membrane and serves as a specific environmental sensor and, in response to environmental changes, auto-phosphorylates and transfers its phosphoryl group to its cognate response regulator. The phosphorylated response regulator, thus activated, binds to a cognate promoter(s) enhancing transcription of a particular gene, or group of genes. Several studies (9–14) have reported the discovery of lead compounds that inhibit kinase activity of TC His kinases. A reevaluation of these findings later noted some of the compounds as acting at other (unspecified) targets (15). In retrospect, the enzymatic activity of His kinases in vivo might be affected by (1) perturbing the cell membrane or, alternatively, (2) the activity of kinase-response regulator pairs might also include cross-talk effects (see Note 1). Assay development based on the interaction between a response regulator and its cognate DNA binding site avoids both mitigating circumstances. We describe a fluorescence polarization assay based on the VanRS TCST system that regulates vancomycin resistance in Enterococcus faecium. Assay development utilizing this system has provided a model system that quantifies the binding between DNA promoter segments and their cognate response regulator protein. Using the VanRS TCST system, we demonstrate inhibition of response regulator protein-promoter segment binding with a known inhibitor. Observed binding constants were comparable to those reported in surface plasmon resonance measurements and gel shift measurements (16). 2. Materials 2.1. Promoter DNA Fragment 1. 5′ -Fluorescein-tagged double-stranded 18-mer oligonucleotide promoter segment H2 based on the vanH promoter studies of Holman et al. (8) was used (see Fig. 1).

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Fig. 1. The vanRS and vanH promoters (R1 and H1 + H2, respectively). Expression of vanH is upregulated by binding of VanR∼P to H1 + H2. VanR also binds to its own promoter R1 regulating its own transcription. The promoter segment used in these studies was based on H2. Coordinates are numbered with respect to both the vanR and vanH transcription start sites as reported by Holman et al. (8). The two individual strands, H2 Forward and H2 Reverse, were chemically synthesized, with H2 Forward arbitrarily selected to carry the 5′ -fluorescein: H2 Forward  5′ Z − TTT TCT TAG GAA ATT AAC H2 Reverse  5′ − GTT AAT TTC CTA AGA AAA where Z = fluorescein. 2. The two single-stranded samples were dissolved in de-ionized water at a concentration of 8 μM and stored at –20 °C. Prior to use, they were annealed by mixing equal volumes of the forward and reverse strands and heated in a thermal cycler at 80 °C for 10 min, followed by slow cooling to room temperature.

2.2. Response Regulator Protein 1. The VanR response regulator protein was overexpressed as a GST-fusion protein in Escherichia coli BL21 DE3 using the full-length vanR sequence inserted between NdeI and HindIII in the multiple cloning site of plasmid pGEX-2TK (Amersham Pharmacia Biotech) as described previously (17). GST-VanR was stored in 1X gel shift buffer (GSB). 2. 1X gel shift buffer (GSB): 20 mM HEPES, pH 7.2, 5 mM MgCl2 , 0.5 mM Ca Cl2 , 0.1 mM Na2 EDTA, and 10% glycerol (see Note 2).

2.3. Compound A 1. The inhibitory action of compound A (2-(2,3,4-trifluorphenyl)-2,3dihydroisothiazole-3-one) (Maybridge Chemical Co., KM-04537) on signal transduction was originally reported by Roychoudhury et al. (18) as part of a search for inhibitors of alginate biosynthesis, a virulence factor in Pseudomonas aeruginosa (see Note 3). 2. Compound A was dissolved at a concentration of 1 mM in water and stored at –20 °C.

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3. Methods 3.1. FP Measurements 1. Binding reactions were run in black 384-well black microtiter plates (Corning). 2. All fluorescence polarization experiments were performed using a Model #1420 fluorometer (Wallac, Turku, Finland) with a measurement time of 1 s per well. Fluorescein was excited at 485 nM, and its emission was measured at 515 nM. Measurements were made at ambient temperature, approximately 25 °C. The microtiter plate was allowed to equilibrate for 30 min at room temperature prior to recording fluorescence polarization measurements. 3. The solutions needed to measure GST-VanR-DNA complex formation were prepared as follows: a. Gel shift buffer (GSB) was prepared as in Section 2.2. b. A dilution series GST-VanR in GSB, 20-μL scale, was prepared by seven successive 3.16-fold dilutions beginning with the highest concentration, 6 μM (see Fig. 2). c. Fluoroscein-labeled H2 DNA was dissolved at 130 nM in GSB.

Fig. 2. Fluorescence polarization (FP) analysis of GST-VanR-PvanH2 complex formation, effect of CpdA. FP was used to measure the concentrations of complexes formed between fluorescein-labeled PvanH2 DNA and GST-VanR. The reaction mixtures contained 13 nM 5′-fluorescein-labeled PvanH2, DNA oligomer, and increasing amounts of GST-VanR, as indicated on the abscissa. The structure of CpdA is inserted in the figure. Three test curves were run using 0, 2.5, and 5.0 μM CpdA. Kd for the control reaction was 30 nM. The addition of 5 μM CpdA increased Kd by a factor greater than 50-fold (see Note 7). (Data shown are based on Erickson et al. (22) with permission of Sage Publications Inc.)

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d. 20, 10, and 6 μM stock solution of CpdA in GSB (see Note 4). 4. Using the solution described in Section 3.1.3, the binding reaction contained, in a volume of 40 μL: a. b. c. d.

GSB sufficient to adjust the total volume to 40 μL. GST-VanR, 20 μL of each dilution in the test series. H2 DNA, 4 μL (see Note 5). Test compound, 4 μL. The highest concentration of CpdA used was 5 μM. Other compounds should be tested at higher concentrations (see Note 6).

5. The G factor was obtained by collecting parallel and perpendicular components of fluorescence from a 40-μL sample containing 8 nM of fluorescein in water with water as the blank. A blank reaction containing 40 μL of 1.0 μM GST-VanR in 1X GSB was used to obtain a correction factor to subtract from the raw-intensity data.

4. Notes 1. Thus far, TCST sensor kinases have been tested as possible drug targets, but studies reported to date have failed to produce anti-infective leads (9–14). TCST kinases may be less suitable targets than their cognate response regulators due to phosphorylation by a promiscuous kinase from another system or to nonspecific phosphorylation from small phosphoryl group donors in the cytosolic fraction. This suggests that pharmacological intervention aimed at response regulators would be more difficult to circumvent. 2. GSB was used in the gel mobility retardation experiments as described by Ulijasz et al. (17) at –80 °C. 3. Compound A was shown previously to inhibit phosphoryl transfer from VanS∼P to VanR by its action on VanR (19). The resultant accumulation of VanS∼P was reversed by supplementing the reaction mixture with additional VanR, but additional VanS had no effect. 4. Compound A inhibits the binding of VanR and VanR∼P to its DNA promoter site. This has been demonstrated by both surface plasmon resonance studies and agarose gel electrophoresis mobility retardation, suggesting a wider scope of action involving the VanR protein DNA binding domain as well (17). The regulatory regions of vanRS and vanHAX contain response-regulator-binding sequences designated R1, H1, and H2, as shown in Fig. 1. 5. The promoter DNA functionally serves as a surrogate ligand. The effect is allosteric, unlike other assays that are crafted on the basis of a competitive interaction at the enzyme active site. 6. As shown in Fig. 2, compound A inhibits the binding of PvanH2 to GST-VanR. The Kd for the control reaction was 30 nM. This result is comparable to promoterresponse regulator interaction values reported by Mattison and Kenney (20) for the OmpR bacterial TCST. It is also comparable to the value of 40 nM reported for the binding of VanR∼P to the vanHAX promoter by gel shift measurement (8), and to the value of approximately 150 nM reported for the binding of GST VanR∼P

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to each of the three promoter segments in the previously reported SPR study (16). Compound A at 1.5, 2.5, and 5.0 μM increased the apparent Kd to 35 nM, 150 nM, and 500 nM. This compares with the four- to five-fold increase reported for the apparent Kd for the GST-VanR∼ P promoter segment interactions reported (16). 7. Z’ is a number calculated according to the formula Z ′ = 1 − 3SD1 + SD2/μ1 − μ2 where SD1 and SD2 are the standard deviations in the experimental and control values, and μ1 and μ2 are the means of the experimental and control values (21). A Z′ value between 0.5 and 1 is considered “an excellent assay.” Assuming a difference of 150 mP units between the two signals, Fig. 2, we can calculate that for Z′ = 0.8, SD1 + SD2 would have to be less than 10.

Acknowledgments We thank F. M. Hoffmann for access to a fluorometer, supported by the University of Wisconsin Clinical Cancer Center. References 1 Hoch, J. A., and Silhavy, T.J. (eds.) (1995) Two-Component Signal Transduction. 1. ASM Press, Washington, DC. 2 Stock, A. M., Robinson, V. L., and Goudreau, P. N. (2000) Two-component signal 2. transduction. Ann. Rev. Biochem. 69, 183–215. 3 West, A. H., and Stock, A. M. (2001) Histidine kinases and response regulator 3. proteins in two component signaling systems. Trends Biochem. Sci. 26, 369–376. 4 Giammarinaro, P., Sicard, M., and Gasc, A. M. (1999) Genetic and physiological 4. studies of the CiaH-CiaR two-component signal-transducing system involved in cefotaxime resistance and competence of Streptococcus pneumoniae. Microbiology 145, 1859–1869. 5 Throup, J. P., Koretke, K. K., Bryant, A. P., Ingraham, K. A., Chalker, A. F., 5. Ge, Y., Marra, A., Wallis, N. G., Brown, J. R., Holmes, D. J., Rosenberg, M., and Burnham, M. K. (2000) A genomic analysis of two-component signal transduction in Streptococcus pneumoniae. Mol. Microbiol. 35, 566–576. 6 Zahrt, T. C., and Deretic, V. (2001) Mycobacterium tuberculosis signal trans6. duction system required for persistent infections. Proc. Natl. Acad. Sci. USA 98, 12706–12711. 7 Kallipolitis, B. H., Ingmer, H., Gahan, C. G., Hill, C., and Sogaard-Andersen, L. 7. (2003) CesRK, a two-component signal transduction system in Listeria monocytogenes, responds to the presence of cell wall-acting antibiotics and affects betalactam resistance. Antimicrob. Agents Chemother. 47, 3421–3429.

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8 Holman, T. R., Wu, Z., Wanner, B. L., and Walsh, C. T. (1994) Identification of the 8. DNA-binding site for the phosphorylated VanR protein required for vancomycin resistance in Enterococcus faecium. Biochemistry 33, 4625–4631. 9 Barrett, J. F., Goldschmidt, R. M., Lawrence, L. E., Foleno, B., Chen, R., 9. Demers, J. P., Johnson, S., Kanojia, R., Fernandez, J., Bernstein, J., Licata, L., Donetz, A., Huang, S., Hlasta, D. J., Macielag, M. J., Ohemeng, K., Frechette, R., Frosco, M. B., Klaubert, D. H., Whiteley, J. M., Wang, L., and Hoch, J. A. (1998) Antibacterial agents that inhibit two-component signal transduction systems. Proc. Natl. Acad. Sci. USA 95, 5317–5322. 10 Barrett, J. F., and Hoch, J. A. (1998) Two-component signal transduction as a 10. target for microbial anti-infective therapy. Antimicrob. Agents Chemother. 42, 1529–1536. 11 Stephenson, K., and Hoch, J. A. (2002) Two-component and phosphorelay signal 11. transduction systems as therapeutic targets. Curr. Opin. Pharmacol. 2, 507–512. 12 Kanojia, R. M., Murray, W., Bernstein, J., Fernandez, J., Foleno, B. D., Krause, H., 12. Lawrence, L., Webb, G., and Barrett, J. F. (1999) 6-oxa isosteres of anacardic acids as potent inhibitors of bacterial histidine protein kinase (HPK)-mediated two-component regulatory systems. Bioorg. Med. Chem. Lett. 9, 2947–2952. 13 Macielag, M. J., Demers, J. P., Fraga-Spano, S. A., Hlasta, D. J., Johnson, S. G., 13. Kanojia, R. M., Russell, R. K., Sui, Z., Weidner-Wells, M. A., Werblood, H., Foleno, B. D., Goldschmidt, R. M., Loeloff, M. J., Webb, G. C., and Barrett, J. F. (1998) Substituted salicylanilides as inhibitors of two-component regulatory systems in bacteria. J. Med. Chem. 41, 2939–2945. 14 Weidner-Wells, M. A., Ohemeng, K. A., Nguyen, V. N., Fraga-Spano, S., 14. Macielag, M. J., Werblood, H. M., Foleno, B. D., Webb, G. C., Barrett, J. F., and Hlasta, D. J. (2001) Amidino benzimidazole inhibitors of bacterial two-component systems. Bioorg. Med. Chem. Lett. 11, 1545–1548. 15 15. Hilliard, J. J., Goldschmidt, R. M., Licata, L., Baum, E. Z., and Bush, K. (1999) Multiple mechanisms of action for inhibitors of histidine protein kinases from bacterial two-component systems. Antimicrob. Agents Chemother. 43, 1693–1699. 16 Smith, E. A., Erickson, M. G., Ulijasz, A. T., Weisblum, B., and Corn, R. M. (2003) 16. Surface plasmon resonance imaging of transcription factor proteins: Interactions of bacterial response regulators with DNA Arrays on gold films. Langmuir 19, 1486–1492. 17 Ulijasz, A. T., Kay, B. K., and Weisblum, B. (2000) Peptide analogues of the 17. VanS catalytic center inhibit VanR binding to its cognate promoter. Biochemistry 39, 11417–11424. 18 18. Roychoudhury, S., Zielinski, N. A., Ninfa, A. J., Allen, N. E., Jungheim, L. N., Nicas, T. I., and Chakrabarty, A. M. (1993) Inhibitors of two-component signal transduction systems: Inhibition of alginate gene activation in Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 90, 965–969.

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19 Ulijasz, A. T., and Weisblum, B. (1999) Dissecting the VanRS signal transduction 19. pathway with specific inhibitors. J. Bacteriol. 181, 627–631. 20 Mattison, K., and Kenney, L. J. (2002) Phosphorylation alters the interaction of 20. the response regulator OmpR with its sensor kinase EnvZ. J. Biol. Chem. 277, 11143–11148. 21 Zhang, J. H., Chung, T. D., and Oldenburg, K. R. (1999) A simple statistical 21. parameter for use in evaluation and validation of high throughput screening assays. J. Biomol. Screen. 4, 67–73. 22 Erickson, M. G., Ulijasz, A. T., and Weisblum, B. (2005) Bacterial 2-component 22. signal transduction systems: A fluorescence polarization screen for response regulator-protein binding. J. Biomol. Screen. 10, 270–274.

18 The Activity of rRNA Resistance Methyltransferases Assessed by MALDI Mass Spectrometry Stephen Douthwaite, Rikke Lind Jensen, and Finn Kirpekar

Summary Resistance to antibiotics that target the bacterial ribosome is often conferred by methylation at specific nucleotides in the rRNA. The nucleotides that become methylated are invariably key sites of antibiotic interaction. The addition of methyl groups to each of these nucleotides is catalyzed by a specific methyltransferase enzyme. The Erm methyltransferases are a clinically prevalent group of enzymes that confer resistance to the therapeutically important macrolide, lincosamide, and streptogramin B (MLSB ) antibiotics. The target for Erm methyltransferases is at nucleotide A2058 in 23S rRNA, and methylation occurs before the rRNA has been assembled into 50S ribosomal particles. Erm methyltransferases occur in a phylogenetically wide range of bacteria and differ in whether they add one or two methyl groups to the A2058 target. The dimethylated rRNA confers a more extensive MLSB resistance phenotype. We describe here a method using matrix-assisted laser desorption/ionization (MALDI) mass spectrometry to determine the location and number of methyl groups added at any site in the rRNA. The method is particularly suited to studying in vitro methylation of RNA transcripts by resistance methyltransferases such as Erm.

Key Words: rRNA methylation; ribosomal antibiotic resistance; RNA mass spectrometry.

1. Introduction Many clinically important antibiotics inhibit the growth of bacteria by blocking protein synthesis on the ribosomes (1–3). These antibiotics bind to regions of the ribosome that are concerned with essential steps in protein synthesis such as peptide bond formation, GTP hydrolysis, and mRNA decoding. The main contact sites for the antibiotics are on the rRNA, rather than on the ribosomal protein components (4), which is consistent with the From: Methods in Molecular Medicine, Vol. 142: New Antibiotic Targets Edited by: W. Scott Champney © Humana Press Inc., Totowa, NJ

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view that the rRNA carries out the primary functions of the ribosome, including formation of the peptide bond (5,6). Not surprisingly therefore, changes in the ribosome structure that confer antibiotic resistance are mainly to be found in the rRNA and consist of nucleotide methylations or base substitutions (4). There are indeed cases of ribosomal protein (r-protein) mutations that confer resistance to ribosome-targeting antibiotics. However, these mutations tend to confer resistance in an indirect manner by influencing the conformation of adjacent rRNA structures that make contact with the antibiotic (7,8). In pathogenic bacteria with multiple rRNA (rrn) operons, resistance to ribosome-targeting drugs is most commonly conferred through modification of the rRNA by specific methyltransferase enzymes (9,10). All the rRNA resistance methyltransferases studied to date use S-adenosyl-L-methionine (AdoMet) as the methyl group donor and contain conserved motifs involved in AdoMet binding (11) and have distinct similarity to Rossmann-fold structures found in proteins that bind other adenosine-based cofactors such as ATP and NAD (12). In other parts of their structures, the rRNA methyltransferases are largely heterogeneous, and these differences presumably enable the enzymes to distinguish their specific target nucleotides. Target nucleotides in 16S rRNA tend to be methylated after assembly of the 30S subunit. The methyltransferases Grm and KamA function in this manner by methylating the assembled 30S subunit at nucleotides G1405 and A1408, respectively, and thereby confer resistance to aminoglycoside antibiotics (13,14) (Escherichia coli rRNA nucleotide numbering is used throughout). These target nucleotides are displayed at the decoding region on the subunit interface sites on the mature 30S subunit (15,16). For methylation to occur, the nucleotides are not only required to be accessible on the surface of the small subunit, but also need to be presented in higher-order structures that are absent in the free 16S rRNA. In contrast, the free 23S rRNA is generally the preferred substrate for methylation prior to its complete assembly with r-proteins to form 50S subunits. Examples include nucleotide G748, the target for the tylosin resistance methyltransferase RlmAII (TlrB) (17); nucleotide A1067, the target for the thiostrepton resistance methyltransferase Tsr (18,19); nucleotide A2058, the target for the MLSB (macrolide, lincosamide, and streptogramin B antibiotic) resistance methyltransferase Erm (20,21); and nucleotides G2470, U2479, and G2535 that are respectively targeted by the orthosomycin resistance methyltransferases EmtA (22), AviRb, and AviRa (23). In addition to methylating the free 23S rRNA substrate, these methyltransferases also specifically recognize their targets within short RNA transcripts, making them ideal for the type of in vitro studies described here.

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The most pervasive of the resistance methyltransferases are those belonging to the Erm family. All Erm methyltransferases specifically methylate the N6 position of nucleotide A2058 in 23S rRNA, but differ as to whether they monomethylate or dimethylate this nucleotide (20,24). Erm monomethyltransferases are found predominantly in drug-producing actinomycetes species and confer the MLSB type I phenotype with high resistance to lincosamides, low to moderate resistance to macrolide and streptogramin B antibiotics (24,25), but no resistance to the latest generation of macrolides, the ketolides (26). Erm dimethyltransferases confer the MLSB type II phenotype with high resistance to all macrolide, lincosamide, and streptogramin B antibiotics (24,27) including ketolides (26). Type II MLSB resistance with dimethylation of the rRNA is the more common mechanism in bacterial pathogens. Nucleotide A2058 is situated in the peptidyl transferase loop of 23S rRNA. This region is inaccessible to Erm methyltransferases in assembled 50S subunit particles, although precursor particles can serve as substrates for ErmC (28). Nucleotide A2058 is methylated by Erm(E) (29) and Erm(S) (30) when displayed in small in vitro RNA transcripts of 26 to 48 nucleotides. However, methylation is much more efficient in slightly larger RNA transcripts that maintain the structure of the peptidyl transferase loop (Fig. 1); such substrates are methylated as efficiently as the intact 23S rRNA and are thus wellsuited for studying Erm methyltransferase activity. Methylation activity can be determined by a number of techniques including the use of radiolabeled AdoMet (30), primer extension with reverse transcriptase (29), and matrixassisted laser desorption/ionization mass spectrometry (MALDI-MS) (17,23). Each method has its own strengths and weaknesses, and one of the advantages with the MALDI-MS approach is the ability to distinguish between mono- and dimethylation at the Erm target nucleotide A2058. Here we describe the preparation of the dimethyltransferase Erm(E) for in vitro activity studies on RNA transcripts (Fig. 1) followed by MALDI-MS analysis of methylation. Detection of singly methylated A2058 by MALDI-MS is demonstrated using the Erm(N) monomethyltransferase (formerly TlrD) (27). 2. Materials 1. Lauria Bertani (LB) broth (32) was used as the rich medium for bacterial cultures. Single-distilled water was used for media, as well as for gels and running buffers; double-distilled water was used for all other buffers and solutions. 2. TMN: 50 mM Tris-HCl, pH 7.8, 10 mM MgCl2 , 100 mM NH4 Cl. 3. Buffer A: 20 mM Tris-HCl, pH 7.5, 10 mM MgCl2 , 1 M NH4 Cl, 10% glycerol, 6 mM -mercaptoethanol. Buffer B: 20 mM Tris-HCl, pH 7.5, 10 mM MgCl2 ,

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Fig. 1. In vitro RNA transcripts representing the peptidyl transferase loop of 23S rRNA. The structures are based on the E. coli 23S rRNA sequence, although the majority of nucleotide positions are highly conserved and thus identical in most bacteria (43). Helices 73, 74, 89, and 90 have been shortened in the in vitro transcripts, and the missing sequences have been replaced with stable tetraloops (except for helix 89). The 3´- and 5´-ends of the structures are positioned in helix 89; this ensures that helix 73 will be stably formed and maintain the structure at the target nucleotide A2058. In structure B, helices 73 and 89 have been shortened further, and the tetraloop capping helix 74 has been altered to give a unique RNase T1 fragment containing the A2058 target (AAAG). Testing with Erm(E) and Erm(N) shows that RNA structures such as these, which contain the complete peptidyl transferase loop and most of helix 73, are methylated as efficiently as intact 23S rRNA.

4.

5.

6. 7. 8. 9.

6 mM -mercaptoethanol. Buffer C: 20 mM Tris-HCl, pH 7.5, 10 mM MgCl2 , 100 mM NH4 Cl, 10% glycerol, 6 mM -mercaptoethanol. Sodium dodecyl sulphate (SDS) gels were used to check the size and purity of the Erm methyltransferase. The stacking gel was 4% and the separation gel 12% polyacrylamide (19:1 acrylamide: bis-acrylamide, electrophoresis grade); protein bands were stained with Coumassie Blue. T7 RNA polymerase (Promega), RNAguard (Amersham), and enzymes for DNA manipulations (New England Biolabs) were used according to the suppliers’ recommendations. Buffers were obtained from the enzyme suppliers. TSC buffer (5X concentrated): 200 mM Tris-HCl, pH 7.9, 30 mM MgCl2 , 50 mM dithiothreitol, 20 mM spermidine. NTP mix: 2.5 mM each of ATP, GTP, UTP, and CTP, pH 7.5. Phenol was distilled and equilibrated with water; stored at –20 °C. Gels for checking RNA transcripts were 10% polyacrylamide (19:1 acrylamide: bis-acrylamide, electrophoresis grade) containing 90 mM Tris-borate, pH 8.3, and 1 mM EDTA. Samples were loaded with 1/5 volume of 10% Ficoll 400 containing 0.05% xylene cyanol and 0.05% bromophenol blue marker dyes. After

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12. 13. 14.

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electrophoresis, gels were stained by soaking for 1 h in 5% acetic acid containing 0.1% toluidine blue, and then destained in 5% acetic acid. RNase A was from Sigma-Aldrich, and RNase T1 was from US Biochemicals. 3-Hydroxypicolinic acid (3-HPA; from Sigma-Aldrich) was dissolved in a 1:1 mixture of HPLC-grade acetonitrile and HPLC-grade water to a concentration of 0.5 M. As well as being used for MALDI matrix preparation, the 3-HPA solution also serves as an excellent RNA denaturing agent for the RNase digestion steps. Poros 50R3 chromatography resin (Applied Biosystems) was suspended in HPLCgrade methanol to approximately 0.2 g/mL. Triethylammonium acetate solution (used at 1 M concentration, pH 7.0) was from Merck. Dowex AG 50W-X8 cation exchange material (acid form) was from BioRad. The material was converted to its ammonium form by five repeated incubations with two volumes of 10 M ammonium acetate, followed by five repeats of washing with two volumes of HPLC-grade water. The cation exchange material was resuspended in approximately two volumes of HPLC-grade water.

3. Methods 3.1. Preparation of Erm(E) Methyltransferase 1. Add 0.5 mL of an overnight culture of E. coli cells harboring a plasmid such as pJEK47 encoding erm(E) (see Notes 1 and 2) to 200 mL of LB broth, containing 100 μg/mL of ampicillin, in a 1 L flask. Shake at 100 rpm in incubator at 37 °C, and measure the optical density (A450 ) every 30 min. Draw a semi-log curve to follow the cell growth. Place the cells on ice when they reach an A450 of 0.4. All the buffers, tubes, and centrifuge rotors should be between 0 °C and 4 °C for the rest of this section. 2. Harvest the cells by centrifugation at 10,000X g for 10 min in a Beckman JA14 rotor, or equivalent. Carefully pour off the supernatant. Wash the cells by resuspending in 200 mL of cold TMN buffer and repeating the centrifugation step. Pour off the supernatant, and resuspend the cells in 20 mL of TMN buffer. Transfer the cell suspension to suitably sized polyethylene tubes (JA20 tubes) on ice. 3. The cell walls are lyzed by sonication, keeping the tube on ice. Wear gloves and ear protection; rinse the sonicator probe with ethanol, and dry before and after use. Sonicate with four bursts at approximately 150 W for 30 s (with a 30-s pause between each burst, or longer if the probe begins to heat up). 4. The cell debris is removed by centrifuging at 30,000X g for 10 min. Transfer the supernatant, containing ribosomes and the Erm(E) methyltransferase, to fresh, cold JA20 tubes. Centrifuge again, to remove any remains of cell walls and cell membranes, and transfer the supernatant to cold Ti50 ultracentrifuge tubes. Fill and balance tubes with cold TMN buffer, and centrifuge at 100,000X g for 3 h at 4 °C in an ultracentrifuge. This, and the following, ultracentrifugation step can be carried out at 20,000X g overnight if the timing is more convenient (see Note 3).

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5. Pour off the supernatant. Keep the tubes on ice while redissolving the ribosome pellets by gentle pipetting in 2.5 mL of buffer A. Allow to stand on ice between 2 and 5 h to wash off the methyltransferase. 6. Centrifuge at 100,000X g for 3 h at 4 °C in Ti50 ultracentrifuge tubes. The ribosomes will pellet leaving the methyltransferase in the supernatant. Collect the supernatant and transfer to a dialysis tube. Seal the tube, excluding air. Dialyze against 200 mL of buffer B in a cold room (4 °C) with stirring; change the buffer every hour (four times in all). 7. Transfer the dialyzed supernatant to Eppendorf tubes. Pellet the methyltransferase by spinning at full speed in an Eppendorf centrifuge (15, 0000 to 20, 000 rpm) for 20 min at 4 °C. Remove the supernatant, keep 15 μL for SDS gel analysis, and discard the rest. Gently redissolve each pellet in 50 μL of buffer C (see Note 3). Take out a total of 15 μL for SDS gel analysis for size (see Note 2) and purity (see Note 4). The rest of the methyltransferase can be stored at –20 °C.

3.2. Transcription of RNA Methylation Substrate 1. The DNA templates for transcription of the RNA methylation substrates (Fig. 1) have been assembled from oligodeoxynucleotides, which have then been cloned into a pGEM plasmid after a T7 RNA polymerase promoter. A restriction site sequence (in this case BamHI) has been incorporated at the 3´-end of the template sequence to facilitate run-off transcription. 2. Double-stranded plasmid DNA containing one of these sequences (Fig. 1) is prepared by standard methods. To 10 μg plasmid DNA in an Eppendorf tube add: 10 μL of 10X Bam HI digestion buffer; H2 O to bring the total volume to 100 μL; and 10 U of BamHI enzyme. Leave overnight at 37 °C in a warm air incubator. 3. Add 200 μL of 0.25 M sodium acetate followed by 750 μL 96% ethanol. Leave in a –20 °C freezer for 1 h. Spin at full speed in an Eppendorf centrifuge (15, 0000 to 20, 0000 rpm) for 20 min at 4 °C. Remove supernatant and wash pellet with 100 μL 70% ethanol. Remove supernatant, dry pellet, and redissolve in 50 μL of H2 O (see Note 5). Check 1 μL of the restricted DNA on a 1% agarose gel alongside 0.1 to 0.2 μg of the uncut DNA. 4. For the T7 transcription reaction, add the following to an Eppendorf tube: 20 μL of 5X TSC buffer; 25 μL of NTP mix; 49 μL of BamHI-cut DNA; 50 U (2.5 μL) of T7 RNA polymerase; and 100 U (3.2 μL) of RNA guard. Incubate at 37 °C for at least 2 h (see Note 6). Stop the transcription by adding 200 μL of H2 O and 300 μL of phenol; vortex 30 s. Recover the RNA by centrifuging the phenol mixture for 1 min and taking the aqueous (upper) phase to a fresh tube. Repeat the extraction procedure with 300 μL of phenol/chloroform (1:1, vol:vol) and then with 300 μL of chloroform. 5. Add 0.1 volume of 2.5 M sodium acetate followed by 2.5 volumes of ethanol to the recovered aqueous phase. Leave in the –20 °C freezer for at least 1 h. Spin at full speed in an Eppendorf centrifuge (15, 0000 to 20, 0000 rpm) for 20 min

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at 4 °C. Remove supernatant and wash the pellet with 100 μL of 70% ethanol. Remove supernatant, dry the pellet, and redissolve in 50 μL of H2 O (see Note 5). The transcript can be checked by running 1 μL on a 10% polyacrylamide gel alongside RNA markers to give a rough estimation of the transcript concentration and size. Stain the gel with toluidine blue after the gel run.

3.3. Methylation of RNA by Erm(E) 1. Each methylation reaction contains 5 μL of the RNA transcript, 24 μL of buffer C containing 1 mM S-adenosylmethionine (AdoMet), and 1 μL of Erm(E). Incubate for 45 min at 30 °C (see Notes 7 and 8). 2. Reactions are stopped by extraction with 1 volume of phenol, then 1 volume of phenol/chloroform, and finally 1 volume of chloroform (as described in Step 4 of Section 3.2). Precipitate the RNA, wash the pellet with ethanol, and redissolve in 6 μL of H2 O as described in Step 5 of Section 3.2. 3. The RNA samples are now ready for analysis by MALDI mass spectrometry (see Note 9).

3.4. Analysis of Methylation by MALDI Mass Spectrometry 1. The methylated RNA substrates are to be digested with RNase A or RNase T1 to produce oligonucleotides of a size suitable for MALDI mass spectrometry analysis (see Note 10). Samples of rRNA (2.5 pmol) are mixed with 0.5 μL of 3-HPA (0.5 M in 50% acetonitrile), 0.1 μg of RNase A, or 10 units of RNase T1 (see Note 11) and H2 O to a final volume of 5 μL and are digested for 2.5 h at 37 °C. 2. Purify the digested fragments on Poros 50R3 columns (see Note 12) as follows: A GelLoader tip (Eppendorf) is flattened at the tip of the extended outlet with forceps, reducing the inner diameter to less than 50 μm. 50 μL of methanol are filled into the GelLoader tip, and 1 μL of the precipitate of Poros 50R3 material in methanol is added. Mount the GelLoader tip onto a 1-mL syringe, and press the methanol through to form a column. The column is washed once with 50 μL of methanol and equilibrated by adding 50 μL of 10 mM triethyl ammonium acetate (TEAA), pH 7.0. Load the RNA digestion in 0.3 M TEAA onto the microcolumn, and wash with 50 μL of 10 mM TEAA and elute with 10 μL of 10 mM TEAA/25% acetonitrile (see Note 13). Dry the eluate and dissolve in water to 1–2 pmol/μL. 3. Prepare samples for MALDI mass spectrometry by mixing on the target 1 μL of the purified RNA digest, 0.5 μL of 3-HPA matrix (see Note 14), and a small volume (∼0.1 μL) of the ammonium-loaded cation exchange material. Leave to air-dry, and remove as much cation exchange material as possible under a microscope using a pipette tip. Sample preparation for MALDI tandem mass spectrometry is performed the same way. 4. The oligonucleotides are analyzed using a Voyager STR MALDI mass spectrometer (Applied Biosystems) in delayed extraction reflector time-of-flight mode detecting

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positive ions (see Note 15). Spectra can be smoothed and calibrated using the Data Explorer software supplied with the mass spectrometer (see Note 16). 5. The exact nucleotide position of a modification can be located by tandem mass spectrometry (23). This can be carried out on a MicroMass MALDI Q-TOF Ultima mass spectrometer in positive ion mode (see Note 17). Generally, the window for parent ion selection is set at 2 m/z units, and the collision energy varies between 40 and 100 eV, depending on the mass of the parent ion. When required, spectra can be smoothed and calibrated using the MassLynx software supplied by the manufacturer.

4. Notes 1. The entire erm(E) gene from Saccharopolyspora erythraea, the producer of the macrolide antibiotic erythromycin, has been cloned into R1-derivative plasmids such as pJEK47 (21) and pJEK48 (33). These plasmids are suitable for expression of large amounts of active Erm(E) in E. coli. 2. With its overall length of 385 amino acids, Erm(E) is significantly longer than most other members of the Erm family. Alignments with other Erm methyltransferases (24,27) show that they are generally 10 to 35 amino acids shorter at their N-termini and approximately 90 amino acids shorter at their C-termini. In one recombinant version of Erm(E), expressed from plasmid pSDdiv (26), we removed the Cterminal 89 amino acids and added a histidine-tag without any apparent loss of methylation activity (see Note 4). 3. The Erm(E) methyltransferase associates with ribosomes under low- to moderatesalt conditions (here, up to 100 mM NH4 Cl) and is released by washing with a high-salt buffer (buffer A with 1 M NH4 Cl) (20,21). The solubility of Erm(E) is reduced in the absence of monovalent ions (buffer B), but the enzyme becomes readily soluble again on increasing the salt concentration (buffer C). 4. Fairly high Erm(E) purity (about 80%) is achieved by this procedure. A higher purity (>95%) can be obtained on Ni-NTA resin (Qiagen) after a histidine-tag has been added to the C-terminal of Erm(E). The activity of Erm(E) and other orthologs we have tried, such as Erm(N), remain unaffected by a C-terminal histidine-tag. However, we observed a reduction in Erm(E) activity with N-terminal tags (34). 5. It is important not to excessively dry nucleic acid pellets, as they can be difficult to redissolve. DNA and RNA pellets are best dried by removing all visible traces of 70% ethanol using a micropipette and then leaving the tubes with the lids open on the bench at ambient temperature for 30 min to allow any remain traces of ethanol to evaporate. 6. Slightly higher yields of in vitro transcripts can be obtained by extending the incubation up to a maximum of 16 h (overnight). Longer times yield diminishing returns, probably due to RNA breakdown. 7. As discussed in the Introduction, members of the Erm family are either mono- or dimethyltransferases, and addition of two methyl groups to the A2058 target by

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the latter type confers the more severe resistance phenotype. All Erm methyltransferases have but a single AdoMet binding site, and thus dimethylation proceeds in a two-step manner (35) requiring recharging of the enzyme with a new cofactor molecule. Under both in vivo and in vitro conditions, Erm(E) is an extremely effective dimethyltransferase adding the second methyl group very rapidly, and it is rare that we find a trace of the monomethylated RNA intermediate. Other Erm dimethyltransferases we have studied, including the streptococcal Erm(B) (36) and the mycobacterial Erm(38) (37), are distinctly less efficient at adding the second methyl group to A2058. The effects of Erm dimethyltransferase inhibitors might first become evident as an accumulation of monomethylated product. In Fig. 2, we have used a recombinant version of the monomethyltransferases Erm(N) (histidinetagged at the C-terminus; see Note 4) to demonstrate MALDI-MS detection of the monomethylated product. Potential methyltransferase inhibitors can be added in buffer C, while maintaining the total volume reaction at 30 μL of buffer C. It will be necessary to stop reactions at a series of time points if IC50 values of potential inhibitors are to be estimated. A rapid and accurate estimate of A2058 dimethylation can be obtained by primer extension with reverse transcriptase (21). Primer extension requires appreciably less RNA than MALDI mass spectrometry (about 10% the amount) but has the disadvantage that it cannot detect monomethylation at the N6 of A2058. Masses of the RNase digestion fragments can be calculated using the Mongo Oligo Mass Calculator (http://www-medlib.med.utah.edu/masspec/mongo.htm). Digestion products smaller than trinucleotides are unsuited for MALDI time-offlight mass spectrometry, because the lower m/z range is crowded with numerous signals including those from the matrix and buffers (31). The RNase T1 digestion products will predominantly harbor 2´-3´ cyclic phosphates under the conditions described here. Increasing the digestion time or enzyme concentration will result in a greater proportion of digestion products with a 3´-phosphate (31); these are heavier by 18.01 Da (monoisotopic mass). Commercial cartridges ready-packed with reverse-phase chromatography material are available from various suppliers including Waters (ZipTip cartridges) and Proxeon (StageTip cartridges). Size fractionation of digestion fragments can be obtained by stepwise elution with increasing concentrations of acetonitrile (38); all the fragments (Figure 2B and 2C) will be eluted by 25% acetonitrile. Other matrices that yield higher sensitivity and/or resolution have been reported for oligonucleotide analysis by MALDI mass spectrometry (39,40). However, we prefer the 3-HPA matrix for this type of application, because it discriminates less between digestion fragments, i.e., nearly all fragments are detected regardless of their nucleotide composition or sequence.

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15. The resolution of the delayed extraction, reflector time-of-flight mass analyzer is required to resolve the approximately 1 Da mass difference between U- and C-nucleotides. The instrument may also be operated in negative ion mode with

(a)

(b) RNase A MH+ fragment 115-117 16-18 53-55 123-125 8-10 22-24 28-30 86-88 58-60 1-3 118-121 129-132 32-35 74-77 38-42 89-93 48-52 94-101 105-114

982.16 999.14 999.14 999.14 1014.15 1014.15 1014.15 1014.15 1015.14 1255.04 1343.21 1344.19 1360.19 1360.19 1656.26 1656.26 1672.26 2675.41 3366.49

Sequence AAC GAU AGU GAU GGC GGC GGC GGC GGU pppGGU AGGC GGAU GGGU GGGU AGAAC AAGAC GAGAC GGAAAGAC GAAAGGGGAU

Fig. 2. (continued)

MALDI-MS Analysis of RNA Methylation (c)

RNase T1 fragments 116-118 17-19 27-29 41-43 79-81 14-16 52-55 37-40 48-51 75-78 83-86 21-25 32-36 67-72 7-13 108-114 100-106 87-95 57-66

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879.17 975.13 975.13 975.13 998.16 999.14 1281.16 1327.21 1327.21 1327.21 1351.22 1586.20 1611.19 1891.24 2219.31 2242.34 2291.34 2853.40 3137.38

AUC-OH UCG CUG UCG ACG AUG UUCG AACG ACAG CAAG AAAG CUUCG UUUAG CCUUCG CACCUCG CACACCG AUAACAG ACCCCUACG UCCCUAUCUG

Fig. 2. (a) MALDI mass spectrum of fragments derived from RNase A digestion of structure A (Fig. 1). The fragment containing the A2058 target nucleotide (GGAAAGAC) runs at m/z 2675.40 (monoisotopic mass; see expanded region in box, and Note 18). Detection of a single methyl group is illustrated by the new signal 14 Da larger that appears in samples treated with the monomethyltransferase Erm(N); the methylation reaction with Erm(N) was stopped before it had run to completion. (b) Theoretical monoisotopic masses of the singly protonated RNase A fragments match the empirical masses to within 0.1 Da (see Note 10). All digestion fragments have 5´-OH and 3´-phosphate unless otherwise noted. The 94-101 fragment (italics) corresponds to the 23S rRNA nucleotides 2056 to 2063 and harbors the Erm target at A2058. (c) Theoretical monoisotopic masses of the singly protonated RNase T1 fragments derived from structure B (Fig. 1). The AAAG fragment at 83-86 (italics) corresponds to 23S rRNA nucleotides 2058 to 2061. This fragment is unique in structure B (but not in structure A) and gives a better resolved and more intense MS peak than the larger RNase A fragment containing nucleotide A2058. similar or even better sensitivity, but instrument stability may be compromised by ion polarity switching if the ion source is contaminated. 16. We first perform an external calibration using appropriate oligodeoxynucleotides; this generally decreases the mass error below 0.5 Da. An internal calibration of the spectrum is subsequently performed using RNase digestion fragments, which are not modified by the methyltransferase. 17. Descriptions of the fragmentation behavior of singly protonated RNA in a MALDI Q-TOF instrument can be found in (41). The predominant sequence ions are of the

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c- and y-type; for the nomenclature of nucleic acid fragment ions, see McLuckey et al. (42). 18. The natural isotopic distribution of 12 C and 13 C (approximately 99:1) leads to multiple signals at 1 Da increments, and this is visible upon closer inspection of the MS peaks. The multiplicity is more pronounced for larger oligonucleotide fragments due to the binomial distribution of the carbon isotopes.

Acknowledgments We thank Birte Vester and Lykke Haastrup Hansen for discussions. Support from the Danish Research Agency (FNU Grants #21-04-0505 and #21-040520), the Nucleic Acid Center of the Danish Grundforskningsfond, and CDC funds are gratefully acknowledged. References 1 Gale, E. F., Cundliffe, E., Reynolds, P. E., Richmond, M. H., and Waring, M. J. 1. (1981) The Molecular Basis of Antibiotic Action. John Wiley and Sons, London. 2 Vázquez, D. (1979) Inhibitors of Protein Biosynthesis. Springer-Verlag, Berlin. 2. 3 Poehlsgaard, J., and Douthwaite, S. (2005) The bacterial ribosome as a target for 3. antibiotics. Nat. Rev. Microbiol. 3, 870–881. 4 Cundliffe, E. (1990) Recognition sites for antibiotics within rRNA. In The 4. Ribosome: Structure, Function and Evolution (W. E. Hill, A. Dahlberg, R. A. Garrett, P. B. Moore, D. Schlessinger, and J. R.Warner, eds.). American Society for Microbiology, Washington DC, pp. 479–490, 5 Green, R., and Noller, H. F. (1997) Ribosomes and translation. Ann. Rev. Biochem. 5. 66, 679–716. 6 Nissen, P., Hansen, J., Ban, N., Moore, P. B., and Steitz, T. A. (2000) The structural 6. basis of ribosome activity in peptide bond synthesis. Science 289, 920–930. 7 Gregory, S. T., and Dahlberg, A. E. (1999) Erythromycin resistance mutations in 7. ribosomal proteins L22 and L4 perturb the higher order structure of 23S ribosomal RNA. J. Mol. Biol. 289, 827–834. 8 Gabashvili, I. S., Gregory, S. T., Valle, M., Grassucci, R., Worbs, M., Wahl, M. C., 8. Dahlberg, A. E., and Frank, J. (2001) The polypeptide tunnel system in the ribosome and its gating in erythromycin resistance mutants of L4 and L22. Mol. Cell 8, 181–188. 9 Vester, B., and Douthwaite, S. (2001) Macrolide resistance conferred by base 9. substitutions in 23S rRNA. Antimicrob. Agents Chemother. 45, 1–12. 10 Douthwaite, S., Fourmy, D., and Yoshizawa, S. (2005) Nucleotide methylations in 10. rRNA that confer resistance to ribosome-targeting antibiotics. In Fine-Tuning of RNA Functions by Modification and Editing (H. Grosjean, ed.), Vol. 12. Springer, New York, pp. 287–309.

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11 Kagan, R. M., and Clarke, S. (1994) Widespread occurrence of three motifs in 11. diverse S-adenosylmethionine-dependent methyltransferases suggests a common structure for these enzymes. Arch. Biochem. Biophys. 310, 417–427. 12 Schubert, H. L., Blumenthal, R. M., and Cheng, X. (2003) Many paths to methyl12. transfer: A chronicle of convergence. Trends Biochem. Sci. 28, 329–335. 13 Skeggs, P. A., Thompson, J., and Cundliffe, E. (1985) Methylation of 16S 13. ribosomal RNA and resistance to aminoglycoside antibiotics in clones of Streptomyces lividans carrying DNA from Streptomyces tenjimariensis. Mol. Gen. Genet. 200, 415–421. 14 Thompson, J., Skeggs, P. A., and Cundliffe, E. (1985) Methylation of 16S 14. ribosomal RNA and resistance to the aminoglycoside antibiotics gentamicin and kanamycin determined by DNA from the gentamicin-producer, Micromonospora purpurea. Mol. Gen. Genet. 201, 168–173. 15 Wimberly, B. T., Brodersen, D. E., Clemons, W. M. J., Morgan-Warren, R. J., 15. Carter, A. P., Vonrhein, C., Hartsch, T., and Ramakrishnan, V. (2000) Structure of the 30S ribosomal subunit. Nature 407, 327–339. 16 Schlünzen, F., Tocilj, A., Zarivach, R., Harms, J., Gluehmann, M., Janell, D., 16. Bashan, A., Bartels, H., Agmon, I., Franceschi, F., and Yonath, A. (2000) Structure of functionally activated small ribosomal subunit at 3.3 Angstroms resolution. Cell 102, 615–623. 17 Liu, M., Kirpekar, F., van Wezel, G. P., and Douthwaite, S. (2000) The tylosin 17. resistance gene tlrB of Streptomyces fradiae encodes a methyltransferase that targets G748 in 23S rRNA. Mol. Microbiol. 37, 811–820. 18 Thompson, J., Schmidt, F., and Cundliffe, E. (1982) Site of action of a 18. ribosomal RNA methylase conferring resistance to thiostrepton. J. Biol. Chem. 257, 7915–7917. 19 Bechthold, A., and Floss, H. G. (1994) Overexpression of the thiostrepton19. resistance gene from Streptomyces azureus in Escherichia coli and characterization of recognition sites of the 23S rRNA A1067 2’-methyltransferase in the guanosine triphosphatase center of 23S ribosomal RNA. Eur. J. Biochem. 224, 431–437. 20 Skinner, R., Cundliffe, E., and Schmidt, F. J. (1983) Site of action of a ribosomal 20. RNA methylase responsible for resistance to erythromycin and other antibiotics. J. Biol. Chem. 258, 12702–12706. 21 Vester, B., and Douthwaite, S. (1994) Domain V of 23S rRNA contains all the 21. structural elements necessary for recognition by the ErmE methyltransferase. J. Bacteriol. 176, 6999–7004. 22 Mann, P. A., Xiong, L., Mankin, A. S., Chau, A. S., Mendrick, C. A., 22. Najarian, D. J., Cramer, C. A., Loebenberg, D., Coates, E., Murgolo, N. J., Aarestrup, F. M., Goering, R. V., Black, T. A., Hare, R. S., and McNicholas, P. M. (2001) EmtA, a rRNA methyltransferase conferring high-level evernimicin resistance. Mol. Microbiol. 41, 1349–1356.

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23 Treede, I., Jakobsen, L., Kirpekar, F., Vester, B., Weitnauer, G. A. B., and 23. Douthwaite, S. (2003) The avilamycin resistance determinants AviRa and AviRb methylate 23S rRNA at the guanosine 2535 base and the uridine 2479 ribose. Mol. Microbiol. 49, 309–318. 24 Weisblum, B. (1995) Erythromycin resistance by ribosome modification. 24. Antimicrob. Agents Chemother. 39, 577–585. 25 Zalacain, M., and Cundliffe, E. (1990) Methylation of 23S ribosomal RNA due to 25. carB, an antibiotic-resistance determinant from the carbomycin producer, Streptomyces thermotolerans. Eur. J. Biochem. 189, 67–72. 26 Liu, M., and Douthwaite, S. (2002) Activity of the ketolide antibiotic telithromycin 26. is refractory to Erm monomethylation of bacterial rRNA. Antimicrob. Agents Chemother. 46, 1629–1633. 27 Roberts, M. C., Sutcliffe, J., Courvalin, P., Jensen, L. B., Rood, J., and Seppälä, 27. H. (1999) Nomenclature for macrolide and macrolide-lincomycin-streptogramin B resistance determinants. Antimicrob. Agents Chemother. 43, 2823–2830. 28 Champney, W. S., Chittum, H. S., and Tober, C. L. (2003) A 50S ribosomal subunit 28. precursor particle is a substrate for the ErmC methyltransferase in Staphylococcus aureus cells. Curr. Microbiol. 46, 453–460. 29 Vester, B., Nielsen, A. K., Hansen, L. H., and Douthwaite, S. (1998) ErmE 29. methyltransferase recognition elements in RNA substrates. J. Mol. Biol. 282, 255–264. 30 Kovalic, D., Giannattasio, R. B., Jin, H. J., and Weisblum, B. (1994) 23S rRNA 30. domain V, a fragment that can be specifically methylated in vitro by the ErmSF (TlrA) methyltransferase. J. Bacteriol. 176, 6992–6998. 31 Kirpekar, F., Douthwaite, S., and Roepstorff, P. (2000) Mapping posttranscrip31. tional modifications in 5S ribosomal RNA by MALDI mass spectrometry. RNA 6, 296–306. 32 Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: 32. A Laboratory Manual. Cold Spring Harbor Press, Cold Spring Harbor, New York. 33 Vester, B., Hansen, L. H., and Douthwaite, S. (1995) The conformation of 23S 33. rRNA nucleotide A2058 determines its recognition by the ErmE methyltransferase. RNA 1, 501–509. 34 Vilsen, I. D., Vester, B., and Douthwaite, S. (1999) ErmE methyltransferase rocog34. nizes features of the primary and secondary structure in a motif within domain V of 23S rRNA. J. Mol. Biol. 286, 365–374. 35 Denoya, C., and Dubnau, D. (1989) Mono- and dimethylating activities and kinetic 35. studies of the ermC 23 S rRNA methyltransferase. J. Biol. Chem. 264, 2615–2624. 36 Douthwaite, S., Jalava, J., and Jakobsen, L. (2005) Ketolide resistance in Strepto36. coccus pyogenes correlates with the degree of rRNA dimethylation by Erm. Mol. Microbiol. 58, 613–622.

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37 Madsen, C. T., Jakobsen, L., and Douthwaite, S. (2005) Mycobacterium smegmatis 37. Erm(38) is a reluctant dimethyltransferase. Antimicrob. Agents Chemother. 49, 3803–3809. 38 Mengel-Jorgensen, J., Jensen, S. S., Rasmussen, A., Poehlsgaard, J., Iversen, J. J., 38. and Kirpekar, F. (2006) Modifications in Thermus thermophilus 23S ribosomal RNA are centered in regions of RNA-RNA contact. J. Biol. Chem. 281, 22108– 22117. 39 Asara, J. M., and Allison, J. (1999) Enhanced detection of oligonucleotides in UV 39. MALDI MS using the tetraamine spermine as a matrix additive. Anal. Chem. 71, 2866–2870. 40 Zhu, Y. F., Chung, C. N., Taranenko, N. I., Allman, S. L., Martin, S. A., 40. Haff, L., and Chen, C. H. (1996) The study of 2,3,4-trihydroxyacetophenone and 2,4,6-trihydroxyacetophenone as matrices for DNA detection in matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Rapid Comm. Mass Spectrom. 10, 383–388. 41 Kirpekar, F., and Krogh, T. N. (2001) RNA fragmentation studied in a matrix41. assisted laser desorption/ionisation tandem quadrupole/orthogonal time-of-flight mass spectrometer. Rapid Comm. Mass Spectrom. 15, 8–14. 42 McLuckey, S. A., Van Berkel, G. J., and Glish, G. L. (1992) Tandem mass 42. spectrometry of small multiply charged oligonucleotides. J. Amer. Soc. Mass Spectrom. 3, 60–70. 43 Cannone, J. J., Subramanian, S., Schnare, M. N., Collett, J. R., D’Souza, L. M., 43. Du, Y., Feng, B., Lin, N., Madabusi, L. V., Müller, K. M., Pande, N., Shang, Z., Yu, N., and Gutell, R. R. (2002) The Comparative RNA Web (CRW) Site: An online database of comparative sequence and structure information for ribosomal, intron, and other RNAs. BMC Bioinformatics 3, 2.

19 Assays for -Lactamase Activity and Inhibition Thammaiah Viswanatha, Laura Marrone, Valerie Goodfellow, and Gary I. Dmitrienko

Summary The ability, either innate or acquired, to produce -lactamases, enzymes capable of hydrolyzing the endocyclic peptide bond in -lactam antibiotics, would appear to be a primary contributor to the ever-increasing incidences of resistance to this class of antibiotics. To date, four distinct classes, A, B, C, and D, of -lactamases have been identified. Of these, enzymes in classes A, C, and D utilize a serine residue as a nucleophile in their catalytic mechanism while class B members are Zn2+ -dependent for their function. Efforts have been and still continue to be made toward the development of potent inhibitors of these enzymes as a means to ensure the efficacy of -lactam antibiotics in clinical medicine. This chapter concerns procedures for the evaluation of the catalytic activity of -lactamases as a means to screen compounds for their inhibitory potency.

Key Words: -lactam antibiotics; -lactamases; antibiotic resistance; chromogenic substrates; -lactamase inhibitors; penicillins; cephalosporins; carbapenems.

1. Introduction -Lactam antibiotics constitute a major class of chemotherapeutic agents currently employed in the treatment of diseases of bacterial origin. This family of antibiotics comprises penicillins and cephalosporins, derived from 6-aminopenicillanic acid (6APA) and 7-aminocephalosporanic acid, respectively, as well as oxacephems, carbapenems, and monobactams. Structures of some of these -lactam antibiotics are shown in Fig. 1. Innate features of oral bioavailability, low toxicity, and high efficacy have contributed to the widespread use of these antibiotics in clinical medicine. From: Methods in Molecular Medicine, Vol. 142: New Antibiotic Targets Edited by: W. Scott Champney © Humana Press Inc., Totowa, NJ

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H N

H S

O

Me O

N

Me

O

H

H N

Cl

S

Me

N

Me

H

Me O

+

S

N

NH2

H OMe H N O

O

HO O

O

NMe2

CO2 Meropenem (Carbapenem)

S

N

H OMe H N O

NH2

N S

CO2

Me N N N N

Moxalactam (7-α-methoxy oxacephem)

N OMe H N O

CO2 CH3

H3C

H S N

N

O Cefepime (Cephalosporin)

S

O

Cefoxitin (Cephamycin)

H2N

O

N

O

CO2

Me CO2

NH2 +

S

N

CO2

Me

O H

Me

Imipenem (Carbapenem)

N

Cloxacillin (Penam)

OH

HN

H

S

O

CO2

Oxacillin (Penam)

OH

H

O

O

CO2

Penicllin G (Penam)

S

N

CO2

Me

N

H2N S

N

O O CH3

HN

Aztreonam (Monobactam)

N O

SO3

Fig. 1. Some clinically important -lactam antibiotics.

Such extensive use, often overuse or abuse, of penicillins and cephalosporins in chemotherapy has led to the development of bacterial resistance to -lactam antibiotics. This phenomenon has been and continues to be the topic of intense investigation in recent years (1–4). In general, microorganisms have been found to gain antibiotic resistance by the use of either one or a combination of the following strategies (5,6): (1) extrusion of the drug by means of an active transmembrane efflux system; (2) acquisition of tolerance to the antibiotic via subtle mutation(s) in the gene encoding for the target protein; (3) production of one or more enzymes capable of deactivating the antibiotic; and (4) rendering the cell less permeable or impermeable to the antibiotic by the loss of porins or channels required for

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its entry. Of these mechanisms, the production of enzymes, viz. -lactamases, happens to be the most commonly encountered and hence the most significant contributor to the emergence of bacterial resistance to -lactam antibiotics (7–9). Furthermore, the promiscuous transfer of the genetic information encoding for the production of -lactamases, not only among the members within a group but also among the diverse genera of microorganisms, via inherently mobile elements such as plasmids or transposons, appears to be a major contributor to the ever-increasing incidences of bacterial resistance to -lactam antibiotics. In light of the above-noted rapid spread of microbial resistance to -lactam antibiotics, strategies to circumvent the adverse action of -lactamase(s) have been and still continue to be developed. These include (1) development of -lactam antibiotic analogues resistant to the action of -lactamase(s), e.g., carbapenems, such as imipenem and meropenem (10), and monobactams, such as aztreonam (11), and (2) production of -lactamase inhibitor(s) which could be co-administered with the -lactam antibiotic in clinical medicine, e.g., clavulanic acid (12), sulbactam (13), and tazobactam (14) (structures given in Fig. 2). According to the currently accepted classification, -lactamases fall into four distinct classes, viz., A, B, C, and D, respectively (15). Of these, the members of classes A, C, and D utilize an active-site serine residue as a nucleophile in their catalytic mechanism involving an acyl-enzyme intermediate akin to that noted in the case of serine proteases (16). The class B -lactamases, in contrast, are metalloproteins with Zn2+ present in their active sites and are often referred to as MBLs (metallo--lactamases). A further division of the MBLs into three subclasses (B1, B2, and B3) has been proposed and is widely accepted (17). The MBLs isolated to date have two distinct Zn2+ binding sites. The class B3 MBLs are carbapenemases that occur in various Aeromonas species. They function with only one zinc ion in the active site and, in fact, appear to be inhibited by zinc binding to the second binding site (18–20). Most

H O

OH

N O

CO2H

calvulanic acid

O

H O2 S

H O2 S

N

N CO2H

sulbactam

O

N

N

CO2H

tazobactam

Fig. 2. Clinically important -lactamase inhibitors.

N

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of the class B1 and class B2 MBLs appear to be capable of functioning as either dizinc or monozinc enzymes, and there has been considerable debate concerning the active-site stoichiometry with respect to zinc under physiological conditions (20). The very interesting and clinically important SPM-1 (Sao Paulo metallo-ß-lactamase) appears to fall somewhere in between the B1 and B2 classifications and is the least effective of the B1 and B2 MBLS at binding the second zinc ion to its active site (21). Active-site metal ion replacement has been achieved under a number of conditions (22,23) and, in some cases (e.g., Zn2+ → Co2+ ), has yielded a catalytically competent enzyme (23). As in the cases of other metallo-proteases, the catalytic mechanism of class B -lactamases does not involve an acyl-enzyme intermediate in the reaction pathway (20,24). 1.1. The Principles Underlying -Lactamase Assay Methods -lactamases catalyze the hydrolysis of penicillins and cephalosporins, as shown in Fig. 3.The assay procedures for the measurement of -lactamase activity are based on the consequences of the hydrolysis of the amide bond of the substrate. Both direct and indirect assay procedures have been developed for the assessment of -lactamase activity. The latter methods exploit some of the innate features of the reaction product; these include (1) its ability to function O O NH

R

S

β-lactamase

N O

R O

H2O

NH S O

+ H

HN CO2

CO2

penicillin (antibiotic)

penicilloic acid (devoid of antibiotic activity) O

O

H N

R

S

H2 O

X β-lactamase

N O

NH

R O

S O

CO2

+ H X

HN CO2

cephalosporin (antibiotic)

cephalosporoic acid (devoid of antibiotic activity)

Fig. 3. Hydrolysis of -lactam antibiotics.

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as an acid by virtue of the newly formed carboxylic acid function (see Fig. 3). This aspect has provided the basis for such assay techniques as (a) alkali-metric titration with the aid of a pH-stat (25), (b) acidimetric analysis involving the use of an indicator such as phenol red or bromocresol purple (26), and (c) manometric assay procedure (27); (2) its ability to function as a reducing agent and thus forming the basis for iodometric analytical procedures (28,29); and (3) the absence of antibiotic activity, the basis for the development of biological assay techniques (30,31). The protocols and the related details of these procedures have been documented (32). The relative merits of some of these methods have been assessed (33). The direct methods, on the other hand, exploit the difference between the substrate and its product with regard to their UV-visible spectra. The lactamase catalyzed reaction is usually monitored by following the change in absorbance at the wavelength where the difference in the molar absorptivity, M , is maximal or near the maximum value. This approach, which was initially employed to monitor the -lactamase catalyzed hydrolysis of cephalosporins (34), has been extended to studies with penicillins as substrates (35,36). The introduction of chromogenic -lactam substrates such as nitrocefin (37), CENTA (38), mCPP (39), PADAC (40), and FAP (6--furylacryloylamidopenicillanic acid) (41) has allowed for spectrophotometric assays to be performed in the visible spectral region, a feature that allows for circum-

H N O

S

H H N

H S N

Me

O

Me

O

H H N O

Nitrocefin

S

H

O

N

O

PADAC

NO2

CO2

H H N

S N

S N

O

CO2

Penicllin G

S

H

NO2

O

H

N

N

CENTA

O O

N

S

O CO2

NH

S

CO2

NO2

CO2

CO2

Depsipeptide Substrate H3C

N

CH3

Fig. 4. Useful substrates for -lactamase assays. mCPP, 1-(3-chlorophenyl)piperazine; PADAC, pyridine-2-azo-p-dimethylanaline cephalosporin.

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venting the problems arising from the high absorbance of most of the substrates in the UV spectral region. The structures of these substrates are shown in Fig. 4. 2. Materials 2.1. Buffers 1. 100 mM of acetate, pH 4.5–5.5. 2. 50 mM of phosphate, pH 6.0–7.5. 3. 50 mM of HEPES (N-(2-hydroxyethyl)piperazine-N´-2-ethane sulfonic acid), pH 6.0–7.5. 4. AMT buffer system (42): 50 mM of acetic acid, 50 mM of MES (2-(N-morpholino)ethane sulfonic acid), and 100 mM of Tris (tris(hydroxymethyl)aminomethane), pH 4.5–9.0 (see Note 1).

2.2. Substrates The chromophoric -lactams, nitrocefin (37) and CENTA (38), are the substrates of choice by virtue of their attractive features that allow for a rapid and reliable spectrophotometric assay procedure for monitoring the activity of -lactamases. The particulars regarding the use of nitrocefin, the most widely used substrate, are provided below: 1. Nitrocefin: A stock solution (2 mM) is prepared by dissolving 10.3 mg of nitrocefin in 0.5 mL of DMSO (dimethyl sulfoxide) and subsequently adding 25 mM HEPES buffer, pH 7.3, or other desired buffer to a final volume of 10 mL. This nitrocefin stock solution (DMSO 5% v/v) is stable for several weeks when stored at 4 °C. 2. CENTA: A stock solution (1 mM) is prepared by dissolving 5.4 mg in 10 mL of 25 mM HEPES, pH 7.3, or other desired buffer. This CENTA stock solution is stable for several days when stored at 4 °C. 3. Benzyl penicillin (Pen G): In instances when Pen G is used as a substrate, a stock solution (10 mM) of this substrate is prepared by dissolving 37 mg of benzyl penicillin (potassium salt) in 10 mL of the desired buffer (see Note 2). Despite the drawback of requiring the use of low-wavelength measurements, assays employing Pen G do have the advantage of low cost and still find significant use in this area.

2.3. Enzyme -lactamase (stock) solutions (0.4–1.0 μM) can be stored at –20 °C without adverse effect on the catalytic activity of the protein (see Note 3). However, they are prone to inactivation when subjected to repeated freezing and thawing.

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To circumvent this problem, it is recommended that the stock solutions of the protein be stored in small (10–20 μL) aliquots and that the remnant of the aliquot be discarded after its use in the desired experiment. -Lactamases are marked with very high catalytic efficiency. Consequently, it is essential to determine, by preliminary experiments, the range in which a linear relationship prevails between the initial rate of reaction and the final concentration of the enzyme in the assay. An enzyme concentration, within a range that allows for an accurate measurement of the initial rate during the progress of the reaction, is to be chosen. 2.3.1. Production and Purification of -Lactamases 1. Purification of IMP-1 as reported (43). An overnight culture of Echerichia coli BL21 (DE3) pLysS transformed with pCIP4 in 5 mL of LB containing 50 μg/mL of kanamycin served as the inoculant for LB medium (500 mL) containing 50 μg/mL of kanamycin, and the culture was incubated at 37 °C, shaking at 200 rpm. At mid-late log phase of culture growth (Abs600 = 0.6–0.8), production of IMP-1 was induced by the introduction of 1 mM IPTG, and growth was allowed to proceed as before for a period of 2–3 h. 2. The cells were collected by centrifugation (6000X g) prior to washing with 50 mM of HEPES buffer, pH 7.3. 3. Cell lysis was achieved by four cycles of freezing and thawing and subsequently treated with DNase and RNase (1 mg of each). Unbroken cells and cell debris were removed by centrifugation (300,000× g), and the supernatant fraction served as a source for IMP-1. 4. Following the addition of ZnSO4 (10 μM), the supernatant was subjected to chromatography on S-Sepharose FF matrix ( Amersham Biosciences) using a 2 × 10-cm column, equilibrated with 50 mM of HEPES buffer, pH 7.3. After washing the column with 3 × 25 mL of equilibrating buffer, proteins bound to the column were eluted with a linear increasing gradient of NaCl in equilibration buffer. 5. Fractions containing -lactamase activity (monitored by nitrocefin hydrolysis) were pooled and dialyzed against 50 mM of HEPES buffer, pH 7.3. 6. The dialyzed material was subsequently applied onto the second cation exchange column, POROS S-20 (Perceptive System) equilibrated with 50 mM of HEPES buffer, pH 7.3. After washing the column with the equilibrating buffer, IMP-1 was eluted with the same buffer containing an increasing concentration of NaCl (0–300 mM). Fractions (5 mL) were collected and assessed for the presence of protein (A280nm ) and -lactamase activity with nitrocefin as substrate. The protein recovered by this procedure was found to be homogeneous by SDS-PAGE. 7. Literature references for the procedures for the production and purification of other representative members of the four classes of -lactamases are summarized in Table 1.

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T. Viswanatha et al. Table 1 Published Procedures for the Isolation and Purification of Some Representative -Lactamases -lactamase Class A B

C D

Enzyme *

Reference

BcI, TEM-1 IMP-1 BcII CcrA VIM-2 L1 P99 Oxa 10

44 45 43 44 46 47 48 49 50

BcI, Bacillus cereus 569H (type I); TEM, class A -lactamases named after the patient (Temoneira) providing the first sample; in the early literature also termed RTEM or R-TEM to emphasize its R plasmid origin; IMP-1, plasmid-encoded class B -lactamases active on imipenem. Pseudomonas aeruginosa 101/477 (most common in Japan); BcII, Bacillus cereus 569H (type II); CcrA, Bacteriodes fragilis TAL636 (Cefoxitin and carbapenem resistant); VIM 1, Verona integron-encoded metallo--lactamase. Pseudomonas aeruginosa VR-143/97 (most common in Taiwan, Verona; L1, Labile enzyme from Stenotrophomonas (Pseudomonas, Xanthomonas) maltophilia; P99, chromosomal -lactamase from Enterobacter cloacae P99; Oxa 10, class D -lactamases active on oxacillin (Acinetobacter species).

2.4. Automated and Semi-automated Procedures for the Assessment of -Lactamase Activity 1. Reagents: 2 mM of nitrocefin, 10.3 mg of nitrocefin is dissolved in 500 μL DMSO and subsequently diluted to 10 mL by the addition of 50 mM of HEPES, pH 7.3. 2. -lactamase: Metallo--lactamase (4 nM) prepared by the dilution of enzyme stock (400 nM) in 50 mM of HEPES, pH 7.3. 3. OXA--lactamase (3–5 nM) of enzyme in 100 mM of phosphate, pH 7.0, containing 25 mM of NaHCO3 . 4. Classes A and C -lactamases (3–5 nM) in 100 mM of phosphate, pH 7.0. 5. 25 mM of HEPES, pH 7.3. 6. 100 mM of phosphate, pH 7.0, containing 25 mM of NaHCO3 (see Note 4). 7. 96-well polystyrene microtitre plates (Costar). 8. Plate reader (Molecular Dynamics, Spectramax 190). 9. Eppendorf multichannel dispenser. 10. Softmax Pro 3.0 for recording rates (total time of recording, 3 min; interval reading, 6 s; and the rate is estimated from the linear increase in absorbance values

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between 20 and 60 s. The total absorbance change is less than 1.0 OD in order to ensure compliance with Beer’s law. 11. Grafit 4.0 for IC50 determination.

3. Methods As noted in Section 1, -lactamases, identified to date, fall into four distinct categories, classes A, B, C, and D. The requirement(s) for the optimal expression of catalytic function, as is to be expected, may depend on the class of -lactamase under investigation. Information concerning such specific requirement(s) is provided in the sections pertaining to the assay procedures for the different types of -lactamases. Given below are the details of items that are common to all the procedures regardless of the class or type of enzymes under study. 3.1. Classes A, C, and D -Lactamases 1. Substrate: 2 mM of nitrocefin stock solution in 50 mM of HEPES, pH 7.3, 10 mM of benzyl penicillin stock in 50 mM HEPES, pH 7.3, or the desired buffer with 25 mM sodium bicarbonate being present in the assays for class D enzymes. 2. Enzyme: -lactamase stock solution (0.4–1.0 μM) diluted with the assay buffer to achieve the desired concentration in the reaction mixture. 3. Spectrophotometric assay procedure: a. Nitrocefin substrate: The enzyme in 950 μL of 50 mM HEPES, pH 7.3, is transferred to a cuvette (10-mm path length) and placed in the sample compartment of the spectrophotometer and allowed to equilibrate at 30 °C for 1 min. The reaction is initiated by the introduction of an aliquot (50 μL) of nitrocefin stock, and the reaction’s progress is monitored by recording the absorbance at 482 nM. The concentrations of nitrocefin and the enzyme, in the reaction mixture of final volume of 1 mL, are 100 μM and 2–4 nM, respectively. b. CENTA substrate: Similar experimental conditions apply when CENTA replaces nitrocefin as the substrate. The reaction can be monitored by following the change in absorbance either at 346 nM (accompanying the hydrolysis of the substrate) or at 405 nM (arising from the elimination of the 2-nitrothiobenzoate chromaphore from the hydrolytic product, cephalosphoroic acid). In the latter procedure based on an increase of absorbance at 405 nM, the reaction should be performed in buffer media of pH >7.0 to ensure the nitro-thiobenzoic acid moiety is in its fully dissociated state. Although CENTA would appear to be a preferable substrate by virtue of its ready solubility in aqueous media (a feature that obviates the need for a cosolvent such as DMSO in the assay), its lower affinity (reflected in high Km -values) for many class B -lactamases (38), relative to that of nitrocefin,

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demands its presence at a relatively high concentration (mM range) to ensure substrate saturation conditions in the assay. c. Benzyl penicillin substrate: The stock solution of the substrate is diluted 10-fold with the buffer (same buffer as in the stock solution) to achieve a final concentration of 1 mM. In a typical assay, 1 mL of the substrate is transferred to each of the two cuvettes, which are subsequently placed in the reference and sample compartments of a double-beam spectrophotometer. Following equilibration at 30 °C for 1 min, the reaction is initiated by the introduction of an aliquot of the enzyme to the substrate in the reference compartment. The reaction’s progress was monitored by recording the increase in absorbance at 232 nm. The final concentration of the substrate and the enzyme in the reaction mixture of total volume of 1 mL are 1 mM and 2–4 nM, respectively.

3.2. Assessment of the Potency and the Mechanism of Action of Inhibitors In the studies with the inhibitors, the procedure is similar to that described above except for the inclusion, in the assay, of the compound to be tested for its potency as an inhibitor of the -lactamase. The protocols to be observed in the experiments designed to evaluate the potency of -lactamase inhibitors have been elegantly addressed in a review (51). These studies are performed under conditions of both with and without prior incubation of the enzyme and the inhibitor. In the latter instance, the inhibitor is introduced into the substrate solution, and the reaction is initiated by the addition of the enzyme. In experiments involving preincubation, the enzyme (0.5–1.0 mL; 0.4–1.0 μM) in an appropriate buffer is treated with the compound (1–10 mM) to be tested for inhibitory potency and allowed to incubate at 30 °C. Appropriate aliquots, drawn at desired intervals, are used to assess the enzyme’s catalytic activity with nitrocefin, benzyl penicillin, or CENTA as substrates, with the final volume of the reaction mixture being 1 mL as outlined above. Estimates of the steady-state kinetic parameters (kcat and KM ) for the hydrolysis of the substrate are achieved by fitting the initial velocity data to the Michaelis–Menten equation with the software package, Grafit 4.0 (Erithacus Software Ltd, Stains, UK). The inhibition constant, Ki , is assessed by Dixon’s procedure (52). A Cornish–Bowden plot (53) of the resulting data is used to establish the mode of inhibition. A M -value of 15,900 M−1 cm−1 at 482 nm for nitrocefin hydrolysis (54) and a M -value of 1,100 M−1 cm−1 for benzyl penicillin hydrolysis (55) are used for the calculation of kcat -values. In the case of CENTA, the following

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M -values are used: –2,500 M−1 cm−1 and +6,400 M−1 cm−1 at 346 nm and 405 nm, respectively (see Note 5). 3.3. Slow-Binding Inhibitors In the case of some inhibitors, when assays are performed by preincubation of the inhibitor with the enzyme, with aliquots being removed for assay as described above, the steady-state rate of hydrolysis may be slow to be attained, as indicated by a lag phase in the rate profile. In each case, the quotient vi /vs (where vi and vs represent the initial and steady-state rates of substrate hydrolysis, respectively) provides a quantitative measure of the lag observed. This situation is encountered in the case of slow-binding inhibitors. The interaction of some thiols with IMP-1 provides an example of this phenomenon (54) (Fig. 5). In these instances, estimates of kinetic parameters are achieved from the steady-state data.

Fig. 5. Thiols as classical and slow-binding inhibitors of IMP-1. Influence of mercaptoacetic acid and benzylmercaptan on the enzymatic activity of IMP-1. All assays were performed in 50 mM Hepes, pH 7.3, at 25 °C with nitrocefin (0.2 mM) as a substrate and IMP-1 (0.8 nM). The progress of nitrocefin hydrolysis was monitored at 482 nm: (1) control with no mercaptan present and (2) in the presence of mercaptoaccetic acid (10 μM). The profile is the same whether the enzyme is preincubated with mercaproacetic acid or exposed to it in the presence of substrate. (3) The enzyme was preincubated with benzylmercaptan (10 μM) for 1 min prior to addition of substrate. (4) The enzyme was exposed to benzylmercaptan (10 μM) in the presence of substrate. (Reprinted from (54) with permission from the American Chemical Society.)

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3.4. Mechanism-Based Inhibitors The class of antagonists referred to as “suicide” or “mechanism-based” reagents is recognized by the enzyme as potential substrates but results in inhibition or inactivation of the enzyme as a consequence of a shift in the normal reaction pathway with the concomitant generation of a highly potent inhibitory species. The characteristics of the “suicide” inhibitors are (1) time dependence of inactivation, (2) the inactivation being proportional to inhibitor concentration at low inhibitor concentrations but independent at high concentrations, and (3) a decrease in the rate of inactivation with an increase in substrate concentration. The latter two kinetic phenomena, saturation kinetics and substrate protection, demonstrate that the inactivation process involves the enzyme’s active site (56). The mechanism of action of the mechanism-based inhibitors of class A -lactamases has been the topic of an elegant review (1). 3.5. Assay Procedure for Mechanism-Based Inhibitors A solution of the enzyme (0.3–1.0 μM) in 0.5 mL of appropriate buffer is incubated at 25 °C with the compound (1.0–10 mM) to be tested for the inhibitory potency to achieve the desired molar excess over that of the enzyme. Aliquots (5–10 μL) of the reaction mixture are removed at various intervals and assayed for enzymatic activity with either nitrocefin or benzyl penicillin, as outlined earlier. An enzyme sample incubated under the conditions mentioned above, but for the exclusion of the inhibitor, is used as a control in these studies. The amount of the inhibitor required to effect total inactivation of the enzyme is estimated by extrapolation of the percent residual activity after prolonged incubation (2–3 h) versus the ratio of the molar excess of the inhibitor over that of the enzyme. The procedures for the assay of class C and class D -lactamases are similar to those used in the case of class A enzymes. The assays of class D enzyme are performed in buffer media containing 25 mM of NaHCO3 because of the need to ensure the maintenance of the essential carbamate group in the active site of the enzyme. 3.6. Class B -Lactamases 1. Reagents: 2 mM of nitrocefin containing stock DMSO (5% v/v), -lactamase (400 nM) containing 100 μM of ZnCl2 , and 100 μg/mL of bovine serum albumin (BSA) (see Note 6). 2. Spectrophotometric assay procedure: An aliquot (950 μL) of -lactamase in 50 mM HEPES, pH 7.3, was introduced to a cuvette (10-mm-pathlength) in the sample compartment of the spectrophotometer and allowed to equilibrate at 30 °C for 1 min.

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The reaction was initiated by the addition of an aliquot (50 μL) of nitrocefin stock. The final concentrations of the substrate, enzyme, and BSA, in a reaction mixture of 1 mL, were 100 μM, 4 nM, and 1 μg/mL, respectively (see Note 6). The progress of the reaction was monitored by recording the increase in the absorbance at 482 nm. Estimates of the steady-state kinetic parameters (KM and kcat ) for the hydrolysis of the substrate are achieved by fitting the initial velocity data to the Michaelis– Menten equation with the aid of Grafit 4.0 (Erithacus Software Ltd, Stains, UK). A M -value of 15,900 M−1 cm−1 at 482 nm for nitrocefin hydrolysis (54) is used for the determination of the kcat -value.

3.7. Inhibition by Chelators Class B -lactamases, like other metalloenzymes, are susceptible to inhibition by chelators. The inhibition can ensue as a consequence of either the interaction of the chelator with the metal ion subsequent to its release from the protein matrix (SN 1 type of mechanism) or while it is still associated with the protein (SN 2-type mechanism) as indicated below (57): E  Zn + C ⇋ E  Zn  C ⇋ E + Zn  C or E∗  Zn  C

where E:Zn represents the holoenzyme, C, the chelator, E:Zn:C, the enzyme:zinc:chelator ternary complex, Zn:C, the zinc:chelator complex, E*:Zn:C, the inactive enzyme:zinc:chelator complex, KI , the concentration of the inhibitor for half-maximal inhibition, and kinact , the first-order rate constant for inactivation. Dixon plots (52), which depict the relationship between the reciprocal activity and the inhibitor concentration, are used to determine the type of inhibition exerted by the chelator. If the inhibition is reversible, the relationship is linear, while it is nonlinear if the process is irreversible. A linear plot of log enzyme activity versus time provides the basis for the determination of t1/2 (time required to effect 50% inhibition of the initial activity) value. Similar experiments are performed at different chelator concentrations and t1/2 value in each case recorded. A plot of t1/2 versus 1/[C], the reciprocal of the chelator concentration according to the procedure of Kitz and Wilson (58) and Silverman (59) is used to estimate Ki and kinact . The interaction of metallo--lactamase, IMP-1, with a variety of commonly used chelators (structures shown below) depicted in Figs. 6 and 7 serves as a typical example of this type of investigation. In these studies, the enzyme, IMP-1, in 50 mM of HEPES, pH 7.3, containing 100 μg/m of BSA was incubated with the chelator at the desired concentration (0–200 mM) for 30 min at 25 °C prior to the determination of catalytic activity with nitrocefin as the substrate (60).

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Fig. 6. Common metal chelators. (a) Structures of DPA (dipicolinic acid); (b) OP (ophenanthroline); (c) PAR (4-(2-pyridylazo)resorcinol); (d) EDTA (ethylenediaminetetraacetic acid); (e) TPEN (N,N,N′ ,N′ -tetrakis(2-pyridylmethyl)ethylenediamine); (f) and Zincon. (Reprinted from (60) with permission from Elsevier.)

Fig. 7. Inactivation of IMP-1 by various Zn2+ chelators. Inactivation of IMP-1 by various Zn2+ chelators. The enzyme (4 nM) in Hepes containing BSA (0.1 mg/ml) was incubated with the chelator at the desired concentration (0–0.2 mM) for 30 min at 25 jC and subsequently assayed for activity with nitrocefin as substrate. In the case of Zincon, the reaction was monitored at 525 nm (instead of 482 nm) in order to minimize interference due to the absorbance of the chelator. For clarity of presentation, inhibition curves (Dixon plots) for DPA and OP are shown in the inset of the figure, while that for PAR was omitted, since it closely resembles the curve obtained with TPEN. •, TPEN; , EDTA; , Zincon. Inset: •, DPA; , OP. (Reprinted from (60) with permission from Elsevier.)

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3.8. Automated and Semi-automated Procedures for the Assessment of -Lactamase Activity The spectrophotometric methods outlined above permit an accurate assessment of the kinetic parameters of the -lactamase catalyzed hydrolysis of its substrates and of the IC50 (concentration of inhibitor needed to effect 50% inhibition of the enzyme’s initial catalytic activity) when an inhibitor is included in the assay. However, these procedures are time-consuming, labor-intensive, and uneconomical in view of the requirement of relatively large amounts of the reaction ingredients in the multiple assays needed to be performed to assess the enzyme’s kinetic parameters. Hence, the introduction of a semi-automated technique or procedure that allows for a rapid and reliable means for the assessment of both the substrate profiles of -lactamase as well as for the evaluation of the potency of its inhibitors is indeed a significant development (61). Given below are the details of an automated assay method, adapted in our laboratory, for the screening of -lactamase inhibitors, using the chromogenic substrate nitrocefin (37). Assays are performed in microtiter plates (of 96 wells) with a final reaction volume of 100 μL. In the procedure used in our laboratory, the eight wells of column 1 (of the microtiter plate) are used for pretreatment of the enzyme prior to its exposure to the substrate. Thus, in the microtiter plate, wells A and B are used for control 1 (no DMSO) and control 2 (with DMSO), and wells C to H are used for incubation of the enzyme with the compound assessed for its inhibitory potency at six different concentrations. The ingredients in control 1, control 2, and test samples are given below (Table 2) (see Note 7). Aliquots (10 μL) of nitrocefin stock are placed in each of the wells in columns 2 and 3 of the microtiter plate. Following incubation for 10 min at 25 °C, 90-μL aliquots from each of the wells in the first column are rapidly transferred with the aid of a multichannel dispenser to the corresponding wells in columns 2 and 3, and the microtiter plate is placed in the Plate Reader. After mixing, the initial rate of the reaction in each well is recorded (at 6-s intervals) between 20 and 60 s at Table 2 Ingredients in Control and Test Reagent

Control 1

Control 2

Buffer: 50 mM HEPES, pH 7.3 DMSO Enzyme Inhibitor (in DMSO)

10 μL — 190 μL —

8 μL 2 μL 190 μL —

Test 8 μL — 190 μL 2 μL

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482 nm. The average of the two determinations (from the reactions in the corresponding wells in the columns 2 and 3 of the microtiter plate) is used to determine percent activity relative to the rate in control 2 (see Note 7). IC50 values can be determined by fitting each inhibition data set to the IC50 -2 parameter equation of Grafit 4.0 (Erithacus Software Ltd., Staines, UK) by nonlinear regression analysis. 3.9. New Directions Although the focus of this article is the discussion of practical and proven methodologies for assaying -lactamase activity, it is worth noting that new approaches continue to be discovered. One very new development involves the Hydrophilic group

O

HN

S

S

N O

Hydrophobic, potential gelation agent

CO2

β−lactamase Hydrophilic group

O

HN

Hydrophobic, Activated gelation agent

+

N O

HS

S

O CO2

Self assembly H2O

Hydrogel formation

Fig. 8. A -lactamase assay based on triggering supramolecular hydrogelation.

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design of -lactamase substrates that, upon hydrolysis of the -lactam bond, release a gelation agent that self-assembles in such a way that water molecules are incorporated into the assembled supramolecular hydrogel scaffold. It is reported that the gelation process is readily visible to the naked eye even in the presence of colored impurities that might interfere with other assay methods (62). Although this method is too new to have been adopted widely in -lactamase research, it represents a very imaginative and promising new direction in this field. Figure 8 illustrates this assay approach. 4. Notes 1. This buffer system provides a constant ionic strength of ∼0.1 over the pH range of 4.5–9.0 and is useful in pH optimum determinations. It is pertinent to note the following limitation(s) of the medium. MES can exert inhibitory action on class B -lactamases, as in the CcrA (63), although no adverse effect was observed in the case of IMP-1 (54). And Tris, the third component of the buffer system, being a good nucleophile at high pH, may interfere in the assay procedures of class A, C, and D -lactamases, which operate via an acylenzyme intermediate in their catalytic mechanism (64). 2. Pen G solutions at pH ≥ 6.0 are prone to undergo hydrolysis even when stored at 4 °C. The hydrolysis of Pen G is accompanied by a M -value of -1100 M−1 cm−1 at 232 nm. The deviation from this value in the assay is a reflection of the breakdown of the substrate during storage. Substrate preparations standing for more than 1 h at 4 °C should be replaced by freshly prepared stock solutions. 3. In the case of metallo--lactamases, stock solutions of the enzyme are prepared in buffer medium (other than phosphate) containing 100 μM of ZnCl2 and 100 μg/mL of BSA to minimize the denaturation of the protein. 4. The pH of the buffer increases upon standing due to loss of the bicarbonate. Buffer is to be replaced by a fresh preparation after 4–6 h. 5. In the case of CENTA as substrate, the reaction can be monitored either at 346 nm or at 405 nm. The increase in the absorbance at 405 nm accompanying the hydrolysis of CENTA is due to the formation of 2-nitrothiobenzoate (NTB− ) anion, an outcome of the propensity of the primary product of the reaction to undergo the relevant elimination event. The M -values of 6,400 M−1 cm−1 (38) and ≈8,000 M−1 cm−1 (65) reported in the studies with CENTA as substrate are considerably lower than the M -values reported (66,67). 6. BSA at the concentration present in the enzyme stock protects the enzyme from denaturation. At the concentration present in the reaction mixture, it does not interfere with the catalytic function of -lactamase but, by its protective action, ensures the reaction’s progress is linear with time. 7. Some DMSO preparations have been found to exert strong inhibitory action even when present at a concentration as low as 1% (v/v) in the reaction mixture. DMSO

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preparations should be checked for this feature, and only those without such adverse effects should be chosen for use in the studies and kept under argon at 4 °C. In these studies, a DMSO preparation with no such adverse effect (the difference between the activity in control 1 and that in control 2 does not exceed 10%) is used.

References 1 Knowles, J. R. (1985) Penicillin resistance—The chemistry of -lactamase 1. inhibition. Acc. Chem. Res. 18, 97–104. 2 Livermore, D. M., and Wooford, N. (2006) The -lactamase threat in Enterobac2. teriacae, Pseudomonas and Acinetobacter. Trends Microbiol. 9, 413–420. 3 Payne, D. J., Du, W., and Bateson, J. H. (2000) -Lactamase epidemiology and 3. the utility of established and novel -lactamase inhibitors. Expert Opin. Invest. Drugs 9, 247–261. 4 Fisher, J. F., Meroueh, S. O., and Mobashery, S. (2005) Bacterial resistance to 4. lactam antibiotics: Compelling opportunism, compelling opportunity. Chem. Rev. 105, 395–424. 5 Walsh, C. (2000) Molecular mechanisms that confer antibacterial drug resistance. 5. Nature 406, 775–781. 6 Martinez, J. L., and Baquero, F. (2000) Mutation frequencies and antibiotic resis6. tance. Antimicrob. Agents Chemother. 44, 1771–1777. 7 Helfand, M. S., and Bonomo, R. A. (2003) -lactamases: A survey of protein 7. diversity. Curr. Drug Targets-Infect. Disord. 3, 9–23. 8 Matagne, A., Dubos, A., Galleni, M., and Frère, J. M. (1999) The -lactamase 8. cycle: A tale of selective pressure and bacterial ingenuity. Nat. Prod. Rep. 16, 1–19. 9 Page, M. I., and Laws, A. P. (1998) The mechanism of catalysis and the inhibition 9. of -lactamases. Chem. Comm. 16, 1609–1617. 10 Rasmussen, B. A., and Bush, K. (1997) Carbapenem-hydrolyzing -lactamases. 10. Antimicrob. Agents Chemother. 41, 223–232. 11 Sykes, R. B., Bonner, D. P., Bush, K., and Georgopapadakou, N. H. (1982) 11. Aztreonam (SQ.26,776) a synthetic monobactam specifically active against Gramnegative bacteria, Antimicrob. Agents Chemother. 21, 85–92. 12 Fisher, J., Charnas, R., and Knowles, J. R. (1978) Kinetic studies on the inacti12. vation of Escherichia coli RTEM -lactamase by clavulanic acid. Biochemistry 17, 2180–2184. 13 English, A. R., Retsema, J. A., Girard, A. E., Lynch, J. E., and Barth, W. E. (1978) 13. CP-45,899, a -lactamase inhibitor that extends the antibacterial spectrum of lactams: Initial bacteriological characterization. Antimicrob. Agents Chemother. 14, 414–419. 14 Micetich, R. G., Maiti, S. N., Spevak, P., Hall, T. W., Yamabe, S., Ishida, N., 14. Tanaka, M., Yamazaki, T., Nakai, A., and Ogawa, K. (1987) Synthesis and

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28 Grove, D. C., and Randall, W. A. (1955) Assay methods of antibiotics. A laboratory 28. manual Medical Encyclopedia, New York, p. 16. 29 Perret, C. J. (1954) Iodometric assay of penicillinase. Nature, 174, 1012–1013. 29. 30 Masuda, G. (1976) Studies on bactericidal activities of -lactam antibiotics on 30. agar plates: The correlation with the antibacterial activities determined by the conventional methods. J. Antibiot. 29, 1237–1240. 31 McGhie, D., Clarke, P. D., Johnson, T., and Hutchison, J. G. P. (1977) Detection 31. of -lactamase activity of Haemophilus influenzae. J. Clin. Path. 30, 585–587. 32 32. Sykes, R. B., and Mathew, M. (1979) Detection and immunology of -lactamases. In -Lactamases (J. M. T. Hamilton-Smith and J. T. Smith, eds.). Academic Press Ltd., London, pp. 17–49. 33 Lucas, T. J. (1979) An evaluation of 12 methods for the demonstration of penicil33. linase. J. Clin. Pathol. 32, 1061–1065. 34 O’Callaghan, C. H., Muggleton, P. W., and Ross, G. W. (1968) Effects of 34. -lactamase from gram-negative organisms on cephalosporins and penicillins. Antimicrob. Agents Chemother. 8, 57–63. 35 Samuni, A. (1975) A direct spectrophotometric assay and determination of 35. Michaelis constants for the -lactamase reaction. Anal. Biochem. 63, 17–26. 36 Waley, S. G. (1974) A spectrophotometric assay of -lactamase action on 36. penicillins. Biochem. J. 139, 789–790. 37 37. O’Callaghan, C. H., Morris, A., Kirby, S., and Shingler, A. H. (1972) Novel method for detection of -lactamases by using a chromogenic cephalosporin substrate. Antimicrob. Agents Chemother. 1, 283–288. 38 Bebrone, C., Moali, C. Mahy, F., Rival, S., Docquier, J. D., Rossolini, G. M., 38. Fastrez, J., Pratt, R. F., Frère, J. M., and Galleni, M. (2001) CENTA as a chromogenic substrate for studying -lactamases. Antimicrob. Agents Chemother. 45, 1868–1871. 39 Perumal, S. K., and Pratt, R. F. (2006) Synthesis and evaluation of ketophosph 39. (on)ates as -lactamase inhibitors. J. Org. Chem. 71, 4778–4785. 40 Jones, R. N., Wilson, H. W., and Novick, W. J. Jr. (1982) In vitro evaluation 40. of pyridine-2-azo-p-dimethylaniline cephalosporin, a new diagnostic chromogenic reagent, and comparison with nitrocefin, cephacetrile, and other -lactam compounds. J. Clin. Microbiol. 15, 677–683. 41 Durkin, J. P., Dmitrienko, G. I., and Viswanatha, T. (1977) N-(2-Furyl) acryloyl 41. penicillin: A novel compound for the spectrophotometric assay of -lactamase I. J. Antibiot. 30, 883–885. 42 Ellis, K. J., and Morrison, J. F. (1982) Buffers of constant ionic strength for 42. studying pH dependent processes. Methods Enzymol. 87, 405–426. 43 Laraki, N., Franceschini,N., Rossolini,G. M., Santucci,P., Meunier, C., de 43. Pauw, E., Amicosante,G., Frère, J. M., and Galleni, M. (1999) Biochemical characterization of the Pseudomonas aeruginosa 101/1477 metallo-ß-lactamase IMP-1 produced by Escherichia coli. Antimicrob. Agents Chemother. 43(4), 902–906.

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44 Kuwabara, S. (1970) Purification and properties of two extracellular -lactamases 44. from Bacillus cereus 569/H. Biochem. J. 118, 457–465. 45 Mathonet, P., Deherve, J., Soumillion, P., and Fastrez, J. (2006) Active TEM-1 45. mutants with random peptides inserted in three contiguous surface loops. Protein Sci. 15, 2323–2334. 46 Toney, J. H., Wu, J. K., Overbye, K. M., Thompson, C. M., and Pompliano, D. L. 46. (1997) High-yield expression, purification, and characterization of active, soluble Bacteroides fragilis metallo-beta-lactamase, CcrA. Protein Expr. Purification 9(3), 355–362. 47 Poirel, L., Naas,T., Nicolas,D., Collet,L., Bellais,S., Cavallo,J.-D., and 47. Nordmann, P. (2000) Characterization of VIM-2, a carbapenem-hydrolyzing metallo-beta-lactamase and its plasmid- and integron-borne gene from a Pseudomonas aeruginosa clinical isolate in France. Antimicrob. Agents Chemother. 44(4), 891–897. 48 Crowder, M. W., Walsh, T. R., Banovic, L., Pettit, M., and Spencer, J. 48. (1998) Overexpression, purification, and characterization of the cloned metallo-lactamase L1 from Stenotrophomonas maltophilia. Antimicrob. Agents Chemother. 42(4), 921–926. 49 Goward, C. R., Stevens, G. B., Hammond, P. M., and Scawen, M. D. (1988) Large49. scale purification of the chromosomal ß-lactamase from Enterobacter cloacae P99. J. Chromat. 457, 317–324. 50 Maveyraud, L., Golemi-Kotra, D., Ishiwata, A., Meroueh, O., Mobashery, S., and 50. Samama, J.-P. (2002) High-resolution X-ray structure of an acyl-enzyme species for the class D OXA-10 -lactamase. J. Amer. Chem. Soc. 124(11), 2461–2465. 51 Bush, K., and Sykes, R. B. (1986) Methodology for the study of -lactamases. 51. Antimicrob. Agents Chemother. 30, 6–10. 52 Dixon, M. (1953) The determination of enzyme inhibitor constants. Biochem. J. 52. 55, 170–171. 53 Cornish-Bowden, A. (1974) A simple graphical method for determining the 53. inhibition constants of mixed, uncompetitive and non-competitive inhibitors. Biochem. J. 137, 143–144. 54 Siemann, S., Clarke, A. J., Viswanatha, T., and Dmitrienko, G. I. (2003) Thiols 54. as classical and slow-binding inhibitors of IMP-1 and other binuclear metallo-lactamases. Biochemistry, 42, 1673–1683. 55 Doucet, N., DeWals, P. Y., and Pelletier, J. N. (2004) Site saturation mutagenesis 55. of Tyr 105 reveals its importance in substrate stabilization and discrimination in TEM-1 -lactamase. J. Biol. Chem. 279, 46295–46303. 56 Abeles, R. H., and Maycock, A. L. (1976) Suicide enzyme inactivators. Acc. Chem. 56. Res. 9, 313–319. 57 Auld, D. S. (1988) Use of chelating agents to inhibit enzymes. Methods Enzymol. 57. 158, 110–114. 58 Kitz, R., and Wilson, I. B. (1962) Esters of methanesufonic acid as irreversible 58. inhibitors of acetylcholinesterase. J. Biol. Chem. 237, 3245–3249.

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59 Silverman, R. B. (2000) The organic chemistry of enzyme catalyzed reactions. 59. Appendix 1 Academic Press, New York, pp. 584–587. 60 Siemann, S., Brewer, D., Clarke, A. J., Dmitrienko, G. I., Lajoie, G., and 60. Viswanatha, T. (2002) IMP-1 metallo--lactamase: Effect of chelators and assessment of metal requirement by electrospray mass spectrometry. Biochim. Biophys. Acta 1571, 190–200. 61 Payne, D. J., Coleman, K., and Cramp, R. (1991) The automated in-vitro 61. assessment of -lactamase inhibitors. J. Antimicrob. Chemother. 28, 775–776. 62 Yang, Z., Ho, P.-L., Liang, G., Chow, K. H., Wang, W., Cao, Y., Guo, Z., and 62. Xu, B. (2006) Using -lactamase to trigger supramolecular hydrogelation. J. Amer. Chem. Soc. ASAP Article; DOI: 10.1021/ja0675604. 63 Fitzgerald, F. M. D., Wu, J. K., and Toney, J. H. Unanticipated inhibition of 63. metallo- -lactamase from Bacterioides fragilis by 4-morpholinoethane sulfonic acid (MES): A crystallographic study at 1.85 À resolution. Biochemistry 37, 6791–6800. 64 Oliver, R. W. A., and Viswanatha, T. (1968) Reaction of tris(hydroxymethyl)64. aminomethane with cinnamoylimidazole and cinnamoyltrypsin. Biochem. Biophys. Acta 156, 422–425. 65 Jones, R. N., Wilson, H. W., Novick, W. J. Jr., Barry, A. L., and Thornsberry, C. 65. (1982) In vitro evaluation of CENTA, a new -lactamase-susceptible chromogenic cephalosporin reagent. J. Clin. Microbiol. 15, 954–958. 66 Ellman, G. L. (1959) Tissue sulfhydryl groups. Arch. Biochem. Biophys. 82, 70–77. 66. 67 Riddles, P. W., Blakely, R. L., and Zerner, B. (1979) Ellman’s reagent: 5,5′ 67. dithiobis-92-nitrobenzoic acid: a reexamination. Anal. Biochem. 94, 75–81.

20 Studies of Enzymes That Cause Resistance to Aminoglycoside Antibiotics Engin H. Serpersu, Can Özen, and Edward Wright

Summary Aminoglycoside antibiotics are highly potent, wide-spectrum bactericidals (1,2). Bacterial resistance to aminoglycosides, however, is a major problem in the clinical use of aminoglycosides. Enzymatic modification of aminoglycosides is the most frequent resistance mode among several resistance mechanisms employed by resistant pathogens (1,3). Three families of aminoglycoside modifying enzymes, O-phosphotransferases, N-acetyltransferases, and N-nucleotidyltransferases, are known to have more than 50 enzymes (1,3,4). In this chapter, determination of enzymatic activity of a single enzyme from each family in the presence and absence of an inhibitor is described.

Key Words: antibiotic resistance; aminoglycosides; aminoglycoside acetyltransferase; aminoglycoside nucleotidyltransferase; aminoglycoside phosphotransferase; enzyme inhibition; drug design.

1. Introduction Aminoglycosides are hydrophilic molecules carrying several hydroxyl and amino functional groups. Most of the aminoglycosides have one or more amino sugars that are attached to a 2-deoxystreptamine (2-DOS) ring. Aminoglycosides that contain 2-DOS are generally further separated into two groups as the neomycin group (4,5-disubstituted 2-DOS) and the kanamycin group (4,6-disubstituted 2-DOS) based on the substitution pattern of the 2-DOS (Fig. 1). Aminoglycoside-modifying enzymes (AGMEs) are promiscuous in From: Methods in Molecular Medicine, Vol. 142: New Antibiotic Targets Edited by: W. Scott Champney © Humana Press Inc., Totowa, NJ

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Fig. 1. Kanamycins (4,6-disubstituted deoxystreptamine, top) and neomycins (4,5disubstituted deoxystreptamine, bottom).

substrate recognition, and many AGMEs show overlapping substrate specificity. Therefore, when a specific inhibitor was developed to inhibit one specific enzyme, it failed to inhibit cell growth because other AGMEs, expressed in multiply resistant bacteria, were not affected (5,6). Similarly, inhibitors based on removal of hydroxyl or amine groups from aminoglycosides may inhibit one specific AGME but usually are substrates of others. Thus, an inhibitor must affect several AGMEs simultaneously to be effective. More recently, various inhibitors based on different strategies have been developed (7–15). Our studies also yielded inhibitors of these three enzymes (16); however, to date, no clinically useful inhibitor has been developed against AGMEs. Due to its importance in testing inhibitors, in this chapter, we will describe activity

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measurements with each class of AGMEs using an example enzyme from each family, an acetyltransferase, a nucleotidyltransferase, and a phosphotransferase. 1.1. Acetyltransferase Activity We will use the aminoglycoside acetyltransferase(3)-IIIb (17) (AAC) (MW: 30 kDa, ∼5 μmoles/min/mg specific activity with kanamycin A as substrate at 24 ºC (17)), which catalyzes the acetylation of aminoglycosides at N3. The reaction is Kanamycin A + Acetyl CoA → 3 − Acetyl − Kanamycin A + CoASH

Activity of this enzyme is measured based on the reduction of 4,4’-Dipyridyl disulfide (18) by CoASH, releasing pyridine-4-thiolate, which can easily be detected at 324 nm spectrophotometrically (19). The reaction is 4 4′ − Dipyridyl disulfide + CoASH → CoA − S − S − pyridine + pyridine − 4 − thiolate

1.2. Nucleotidyltransferase Activity Aminoglycoside nucleotidyltransferase(2˝)-Ia (21) (ANT) (MW: 25 kDa, specific activity for tobramycin nucleotidylation: 0.4 units/mg at 30 °C using 40 μM tobramycin and 1.0 mM MgATP (22) is one of the most prevalent enzymes causing resistance to aminoglycosides (23) and catalyzes the covalent attachment of adenosine monophosphate (AMP) to the 2˝ position of 4,6disubstituted aminoglycoside antibiotics (21,24). The overall reaction is MgATP + 2 − hydroxyaminoglycoside → MgPPi + AMP − 2 − aminoglycoside

A discontinuous coupled enzyme assay based on the conversion of one molecule of the reaction product, inorganic pyrophosphate, to two molecules of inorganic phosphate was utilized to measure the steady-state rate of the reaction catalyzed by ANT (21,22,25). Inorganic pyrophosphatase catalyzes hydrolysis of pyrophosphate to inorganic phosphate as shown below, which is then determined spectrophotometrically (25,26): MgPPi →Mg2+ + 2Pi 

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1.3. Phosphotransferase Activity The enzyme used in these studies is the aminoglycoside phosphotransferase(3’)IIIa (28) (APH) (MW: 31 kDa, specific activity for kanamycin A phosphorylation: 2.0 units/mg at 24 °C (29,30)). APH catalyzes phosphorylation of the 3′ -OH position on aminoglycosides. The enzyme is purified based on McKay et al. (28), with modifications described in Özen and Serpersu (30). The catalytic reaction is Mg2+ + ATP + KanA → Mg2+ + ADP + KanA − PO− 3

The rate of this reaction is determined by coupling it to the well-known pyruvate kinase-lactic dehydrogenase coupled enzyme system where phosphoenolpyruvate (PEP) is converted to pyruvate, which is then reduced to lactate while NADH is oxidized to NAD+ : PEP + Mg2+ + ADP − PyruvateKinase → Pyruvate + Mg2+ + ATP

Pyruvate + NADH —-(Lactate Dehydrogenase)→ Lactate + NAD+ . 2. Materials 2.1. Acetyltransferase Activity 1. 2. 3. 4. 5. 6. 7.

Spectrophotometer. 0.5 M Tris-HCl buffer, pH 7.5. Store at 4 °C. 0.25 M EDTA. Store at 4 °C. 100 mM AldrithiolTM -4 (4,4’-dipyridyl disulfide). Store at –80 °C. 50 mM acetyl coenzyme A. Store at –80 °C. 5 mM kanamycin A. Store at –20 °C or –80 °C. AAC sample.

2.2. Nucleotidyltransferase Activity 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

Spectrophotometer. 0.25 M HEPES buffer, pH 7.5. Store at 4 °C. 1.0 M KCl. Store at 4 °C. 100 mM MgCl2 . Store at 4 °C. 2.5 mM tobramycin. Store at –20 °C. 20 mM ATP. Prepare in ice, and adjust pH to 5.0–6.0. Store at –20 °C. Inorganic pyrophosphatase (follow manufacturer’s storage instructions). 1.0% (w/v) Ammonium molybdate in H2 O. Store at room temperature. 10% (w/v) SDS in acetic acid/acetate, pH 4.0. Store at room temperature. 10% Ascorbic acid in ddH2 O. Prepare immediately before use. 0.5–1.0 mg/mL of ANT.

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2.3. Phosphotransferase Activity 1. 2. 3. 4. 5. 6. 7.

Spectrophotometer. 0.5 M TrisHCl buffer, pH 7.5. Store at 4 °C. 1.0 M KCl. Store at 4 °C. 100 mM MgCl2 . Store at 4 °C. 50 mM kanamycin A. Store at –20 °C. 20 mM ATP. Prepare in ice, and adjust pH to 5.0–6.0. Store at –20 °C. 10 mM phospho(enol)pyruvate (PEP). Prepare in 1.0 mM TrisHCl, pH 7.5. Store at –20 °C. 8. 10 mM NADH. Prepare in 1.0 mM TrisHCl, pH 7.5. Store at –20 °C. 9. Pyruvate kinase (PK) and L-lactate dehydrogenase (LDH) mixture, each 750 units/mL. Store at 4 °C. 10. APH sample. In order to obtain a measurable rate, adjust the concentration to 100 μM.

3. Methods 3.1. Acetyltransferase Activity 1. Turn on the spectrophotometer, and set the wavelength to 324 nm and the run time to several minutes. 2. Prepare the assay mix by adding 856 μL distilled water (resistivity 17.5 M-cm or better), 100 μL 0.5 M TrisHCl, and 4 μL 0.25 M EDTA into a quartz cuvette. 3. Equilibrate the mix to the temperature at which the assay will be conducted. 4. Add 2 μL 50 mM acetyl coenzyme A, 8 μL 100 mM 4,4’-dipyridyl disulfide, and 10 μL 100 μM (∼3 mg/mL) AAC sample and mix (see Notes 1 and 2). 5. Blank the spectrophotometer using the mixture prepared in Step 4. 6. Start the run and determine the background rate ( absorbance units/min) for pyridine-4-thiolate formation. 7. Add 20 μL 5 mM kanamycin A, quickly mix, and continue the run. 8. After the run is complete, calculate the rate of increase ( absorbance units/min) in pyridine-4-thiolate absorption using a linear part of the plot. Subtract the background rate from this value to find the enzymatically catalyzed reaction rate (see Notes 3 and 4). 9. Using the rate calculated in Step 8 and the molar extinction coefficient of pyridine4-thiolate (19,800 M−1 cm−1 at 324 nm), calculate the enzyme activity in units (one unit is defined as μmoles of substrate utilized per min). 10. Calculate the specific enzyme activity (μmoles/min/mg or units/mg) by dividing the activity determined in Step 9 by the amount (mg) of AAC in the assay cuvette. AAC activity is ∼5 units/mg (17) (see Notes 5 and 6).

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3.2. Nucleotidyltransferase Activity 1. Turn on the spectrophotometer, and set the wavelength to 820 nm. Prepare a 30 °C water bath. 2. Aliquot 700 μL of 10% (w/v) SDS in acetic acid/acetate, pH 4.0 (stopping solution), in a test tube for each time point of the assay. 3. Prepare 1:1 mixture of 1.0% (w/v) ammonium molybdate:10% ascorbic acid. 4. Prepare the assay mix by adding 280 μL of distilled water (resistivity 17.5 M-cm or better), 25 μL of 0.25 M HEPES, 50 μL of 1.0 M KCl, 75 μL of 100 mM MgCl2 , 25 μL of 20 mM ATP, and 2.0 units (5 μL of 400 units/ml solution) of inorganic pyrophosphatase and 25 μL of ANT into a plastic cuvette (see Note 7). 5. Equilibrate the mix to 30 °C in a water bath. 6. Start reaction by addition of 20 μL of tobramycin. Run a parallel blank reaction by adding water instead of tobramycin (see Notes 8 and 9). 7. Every 2 min remove 100 μL from each reaction mixture and add to tube containing stopping solution. 8. After all time points are complete (3 or 4 for 500 μL assay volume), add 200 μL ammonium molybdate:ascorbic acid solution. Let stand 30 min at room temperature. 9. Read each tube at 820 nm using tubes without tobramycin from each time point as a blank (see Note 10). 10. Calculate rate by plotting A820 vs. time. A reading of 1.0 at 820 nm corresponds to 260 nanomoles of Pi formed. Therefore, a reading of 1.0 results from the formation of 130 nanomoles of MgPP i (and also TobAMP) in the initial reaction. Calculate the specific enzyme activity (units/mg) by dividing the rate by the amount (mg) of ANT in the assay. 11. When present, inhibitors must be added at Step 4 above (see Notes 11 and 12). 12. As indicated earlier, for inhibitors with low water solubility, DMSO, ethylene glycol, or hexylene glycol can be used in activity assays up to a few percent without affecting the enzyme activity (16,20).

3.3. Phosphotransferase Activity 1. Turn on the spectrophotometer, and set the wavelength to 340 nm and the run time to 2.0 min. 2. Prepare the assay mix by adding 495 μL of distilled water (resistivity 17.5 MO-cm or better), 100 μL of 0.5 M TrisHCl, 100 μL of 1.0 M KCl, 15 μL of 100 mM MgCl2 , 10 μL of 50 mM aminoglycoside stock, 50 μL of 20 mM ATP, and 200 μL of 10 mM PEP into a cuvette with 1-cm path length (see Note 13). 3. Equilibrate the mix to the temperature at which the assay will be conducted. 4. Blank the spectrophotometer using the assay mix (see Note 14). 5. Add 15 μL of 10 mM NADH and 5.0 μL of PK/LDH into the cuvette and mix well (see Note 15).

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6. Start the run. Observe a flat baseline for the initial 10 to 20 s, add 10 μL of 100 μM APH, quickly mix, and continue the run. 7. After the run is complete, calculate the rate of decrease ( absorbance units per minute) in NADH absorption using a linear stretch of the plot (see Note 16). 8. Using the rate calculated in Step 7 and the molar extinction coefficient of NADH (6.22 × 103 M−1 cm−1 at 340 nm), calculate the enzyme activity in units (a unit is defined as one μmole/min) (see Note 17). 9. Calculate the specific enzyme activity (units/mg) by dividing the activity determined in Step 8 by the amount (mg) of APH in the assay cuvette (see Notes 18–20).

4. Notes 1. Keep 4,4’-dipyridyl disulfide, acetyl coenzyme A, kanamycin A stocks, and AAC sample on ice. 2. If multiple assays are to be conducted, one can prepare a stock solution of the assay mix by multiplying the amounts given at Step 2 of the Methods section with the number of assays. This solution can be stored on ice for several hours. Use 962 μL from the prepared stock for each assay. 3. In order to ensure that no other assay component except AAC is the limiting factor in the assay, double the amount of AAC added in another run and confirm the doubling of the rate. 4. Mercaptoethanol (BME) can be used as a positive control to test the assay mix. Add 5 μL of 5 mM BME into the cuvette. An immediate increase in absorbance should be observed. 5. When present, inhibitors should be added in Step 4 before the baseline determination. 6. Inhibitors with low water solubility can be added from stocks dissolved in DMSO, ethylene glycol, or hexylene glycol. A matching concentration of solvent needs to be present in assays performed without the inhibitor. Generally, AGMEs are tolerant to the presence of these solvents in the assay medium up to several percentage points. 7. The assay can be scaled up to 1.0 mL if more time points are required. 8. Initial tests to determine the limits of linearity of the assay vs. enzyme concentration (Fig. 2) and linearity of activity vs. time (Fig. 3) are strongly recommended. 9. If desired, commercial preparations of inorganic pyrophosphate can be used in place of tobramycin to construct a standard curve to determine absorbance vs. inorganic phosphate for a particular spectrophotometer. This method also can confirm the activity of the inorganic pyrophosphatase is not a limiting factor in the assay. 10. If multiple assays are to be conducted, prepare an assay mix stock that can be stored at 4 °C for a couple of hours by multiplying the amounts given at Step 2 of the Methods section with the number of assays.

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Fig. 2. A typical experiment showing the linearity of the assay as a function of ANT concentration. Incubation time was 5 min. 11. ANT is an unstable enzyme. It should be kept on ice at all times and should be used within 48 h of preparation. ANT should not be frozen. 12. Substrate inhibition is observed with ANT (22,27). This phenomenon must be considered when designing detailed kinetic analysis using this enzyme.

Fig. 3. Accumulation of inorganic phosphate as a function of assay time. ANT concentration was 0.04 mg/mL. Data represent the average of three measurements with different batches of enzyme. Errors smaller than the size of data points are not visible in early time points.

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13. Keep stock solutions of kanamycin A, ATP, PEP, NADH, PK/LDH, and APH on ice. 14. If multiple assays are to be conducted, one can prepare an assay mix stock that can be stored at 4 °C for a couple of hours by multiplying the amounts given at Step 2 of the Methods section with the number of assays. Use 970 μL from the prepared stock for each assay. 15. NADH stocks will undergo oxidation and take a yellowish color within a couple of weeks. The degree of oxidation can be estimated from the absorbance reading when 15 μL of NADH are added into the assay mix (100% NADH should give an absorbance reading of 0.95). Thus, small stocks of NADH should be prepared to be consumed in a short period of time. Also, repeated freeze/thaw cycles of NADH solutions are not recommended. 16. In order to ensure that no other assay component except APH is the limiting factor for the measured rate, add another 10 μL of APH after a linear decrease in absorbance is observed. The rate should double after the second addition. 17. ADP can be used as a positive control to check the assay mix components in case no activity is observed. Add 10 μL of 10 mM ADP to observe approximately a 0.6-unit decrease in absorbance. 18. In these types of assays, coupling enzyme(s) should not create a bottleneck for observed activity. To guarantee this, activity units per unit volume for coupling enzymes should be at least 10-fold higher than the one for the enzyme studied. 19. Substrate inhibition is observed with APH at high concentrations, which must be considered in detailed kinetic experiments. 20. APH activity is not affected by DMSO, ethylene glycol, or hexylene glycol up to several percent presence of these solvents, which can be used in assays of inhibitors with low solubility in water (16,20).

References 1 Umezawa, H. (1974) Biochemical mechanism of resistance to aminoglycosidic 1. antibiotics. Adv. Carbohydrate Chem. Biochem. 30, 183–225. 2 Davies, J. E. (1991) In Antibiotics in Laboratory Medicine (V. Lorian, ed.). 2. Williams and Wilkins, Baltimore, MD, pp. 691–713. 3 Davies, J. E. (1994) Inactivation of antibiotics and the dissemination of resistance 3. genes. Science 264, 375–382. 4 4. Shaw, K. J., Rather, P. N., Hare, R. S., and Miller, G. H. (1993) Moleculargenetics of aminoglycoside resistance genes and familial relationships of the aminoglycoside- modifying enzymes. Microbiol. Rev. 57, 138–163. 5 Allen, N. E., Alborn, W. E., Hobbs, J. N., and Kirst, H. A. (1982) 75. Hydroxytropolone—An inhibitor of aminoglycoside-2”-O-adenylyltransferase. Antimicrob. Agents Chemother. 22, 824–831.

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6 Williams, J. W., and Northrop, D. B. (1979) Synthesis of a tight-binding, multi6. substrate analog inhibitor of gentamycin acetyltransferase I. J. Antibiot. 32, 1147–1154. 7 Yu, L. P., Oost, T. K., Schkeryantz, J. M., Yang, J. G., Janowick, D., and 7. Fesik, S. W. (2003) Discovery of aminoglycoside mimetics by NMR-based screening of Escherichia coli A-site RNA. J. Amer. Chem. Soc. 125, 4444–4450. 8 Agnelli, T., Sucheck, S. J., Marby, K. A., Rabuka, D., Yao, S. L., Sears, P. S., 8. Liang, F. S., and Wong, C. H. (2004) Dimeric aminoglycosides as antibiotics. Angew. Chem. Intl. Ed. 43, 1562–1566. 9 Sucheck, S. J., Greenberg, W. A., Tolbert, T. J., and Wong, C.-H. (2000) Design of 9. small molecules that recognize RNA: Development of aminoglycosides as potential antitumor agents that target oncogenic RNA sequences. Angew. Chem. Intl. Ed. 39, 1080–1084. 10 Asensio, J. L., Hidalgo, A., Bastida, A., Torrado, M., Corzana, F., Chiara, J. 10. L., Garcia-Junceda, E., Canada, J., and Jimenez-Barbero, J. (2005) A simple structural-based approach to prevent aminoglycoside inactivation by bacterial defense proteins. Conformational restriction provides effective protection against neomycin-B nucleotidylation by ANT4. J. Amer. Chem. Soc. 127, 8278–8279. 11 Griffey, R. H., Hofstadler, S. A., and Swayze, E. E. (2003) Preparation of 11. heterocyclic 2-deoxystreptamine aminoglycoside analogues and characterization of their interaction with RNAs by use of electrospray ionization mass spectrometry. Carbohydrate-Based Drug Discovery 2, 483–499. 12 Haddad, J., Kotra, L. P., Llano-Sotelo, B., Kim, C., Azucena, E. F., Liu, M. Z., 12. Vakulenko, S. B., Chow, C. S., and Mobashery, S. (2002) Design of novel antibiotics that bind to the ribosomal acyltransfer site. J. Amer. Chem. Soc. 124, 3229–3237. 13 13. Fridman, M., Belakhov, V., Yaron, S., and Baasov, T. (2003) A new class of branched aminoglycosides: Pseudo-pentasaccharide derivatives of neomycin B. Org. Lett. 5, 3575–3578. 14 Boehr, D. D., Draker, K. A., Koteva, K., Bains, M., Hancock, R. E., and 14. Wright, G. D. (2003) Broad-spectrum peptide inhibitors of aminoglycoside antibotic resistance enzymes. Chem. Biol. 10, 189–196. 15 He, Y., Yang, J., Wu, B., Robinson, D., Sprankle, K., Kung, P.-P., Lowery, K., 15. Mohan, V., Hofstadler, S., Swayze, E. E., and Griffey, R. (2004) Synthesis and evaluation of novel bacterial rRNA-binding benzimidazoles by mass spectrometry. Bioorg. Med. Chem. Lett. 14, 695–699. 16 Welch, K. T., Virga, K. G., Brown, C. L., Wright, E., Lee R. E., and Serpersu, E. H. 16. (2005) Inhibitors of aminoglycoside-modifying enzymes based on the 1,3-diamine motif of the 2-deoxystreptamine ring. Bioorg. Med. Chem. 13, 6252–6263. 17 Owston, M. A., and Serpersu, E. H. (2002) Cloning, overexpression, and 17. purification of aminoglycoside antibiotic 3-acetyltransferase-IIIb: Conformational studies with bound substrates. Biochemistry 41, 10764–10770.

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18 Williams, J. W., and Northrop, D. B. (1978) Kinetic mechanisms of gentamycin 18. acetyltransferase-I—Antibiotic-dependent shift from rapid to nonrapid equilibrium random mechanisms. J. Biol. Chem. 253, 5902–5907. 19 Grassetti, D. R., and Murray, J. F. (1967) Determination of sulfhydryl groups with 19. 2,2’- or 4,4’-dithiodipyridine. Arch. Biochem. Biophys. 119, 41–49. 20 Serpersu, E., and Dan, J. (2004) Effects of solvents on the activity and substrate 20. preference of the aminoglycoside-3-acetyltransferase-IIIb. Biophys. J. 86, 93a–93a. 21 Wright, E., and Serpersu, E. H. (2004) Isolation of aminoglycoside 21. nucleotidyltransferase(2”)-Ia from inclusion bodies as active, monomeric enzyme. Protein Expr. Purif. 35, 373–380. 22 Wright, E., and Serpersu, E. H. (2005) Enzyme-substrate interactions with 22. an antibiotic resistance enzyme: Aminoglycoside nucleotidyltransferase(2”)Ia characterized by kinetic and thermodynamic methods. Biochemistry 44, 11581–11591. 23 Shimizu, K., Kumada, T., and Hsieh, W. C. (1985) Comparison of aminoglycoside 23. resistance patterns in Japan, Formosa, and Korea, Chile, and the United States. Antimicrob. Agents Chemother. 27, 282–288. 24 Gates, C. A., and Northrop, D. B. (1988) Substrate specificities and structure 24. activity relationships for the nucleotidylation of antibiotics catalyzed by aminoglycoside nucleotidyltransferase-2”-I. Biochemistry 27, 3820–3825. 25 Ekman, D. R., DiGiammarino, E. L., Wright, E., Witter, E. D., and Serpersu, E. H. 25. (2001) Cloning, overexpression, and purification of aminoglycoside antibiotic nucleotidyltransferase (2 ‘)-Ia: Conformational studies with bound substrates. Biochemistry 40, 7017–7024. 26 Ames, B. N. (1965) Assay for inorganic phosphate. Meth. Enzymol. 7, 115–118. 26. 27 Gates, C. A., and Northrop, D. B. (1988) Alternative substrate and inhibition27. kinetics of aminoglycoside nucleotidyltransferase-2”-I in support of a TheorellChance kinetic mechanism. Biochemistry 27, 3826–3833. 28 McKay, G., A., Thompson, P. R., and Wright, G. D. (1994) Broad spectrum 28. aminoglycoside phosphotransferase type III from Enterococcus: Overexpression, purification and substrate specificity. Biochemistry 33, 6936–6944. 29 Ozen, C., Malek, J. M., and Serpersu, E. H. (2006) Dissection of aminoglycoside29. enzyme interactions: A calorimetric and NMR study of noemycin B binding to the aminoglycoside phosphotransferase(3’)-IIIa. J. Amer. Chem. Soc. (in press). 30 Özen, C., and Serpersu, E. H. (2004) Thermodynamics of aminoglycoside binding 30. to aminoglycoside-3’-phosphotransferase IIIb studied by isothermal titration calorimetry. Biochemistry 43, 14667–14675.

Index A Agarose gel electrophoresis, 13, 15, 17, 19 Aminoglycosides, 263–264 Aminoglycoside modifying enzyme assays, aminoglycoside acetyltransferase assay, 265 aminoglycoside nucleotidyltransferase assay, 266 aminoglycoside phosphotransferase assay, 266–267 Antibiotic mechanisms of inhibition, 2 Antibiotic target identification, 3 B Bacillus subtilis, 26 Bacterial colony counting, 58, 66 Bacterial efflux pump inhibitors (EPI), 187–188 Bacterial cell extracts, 28, 42, 57, 67, 78, 98, 110, 123, 227, 246 Bacterial growth media, broth growth media, 30, 38, 56, 64, 68, 120, 189, 225 minimal media, 55–56, 64, 77, 93, 120 Bacterial membrane preparations, 134, 136 Beta-galactosidase assay, 80 Beta-keto acyl carrier protein synthase III (FabH), 207 Beta-lactamase, 241–244 Beta-lactamase assays, 244–245, 250–251, 252–256 Beta-lactamase isolation, 246 BLASTP, 5 Bocillin, 139 Broth growth media, 30, 38, 56, 64, 68, 120, 189, 225 C Calcein leakage assay, 165 Calf thymus DNA activation, 30 Cationic antimicrobial peptides, 155–156 Cell-free translation assays, 88 bacterial translation, 100 human cell translation, 102 yeast cell translation, 101

Chaperone proteins, 75 Circular dichroism assay, 168 D Database of Essential Genes (DEG), 5 Decatenation assay for topo IV, 15 DNA cleavage assay, 17–19 DNA gyrase, 11–12 DNA polymerase III, 26 DNA polmyerase III assay, 28, 32 DNA polymerase III, isolation 27–30 DNA supercoling assay, 13–15 DNA topoisomerase IV, 11–12 DnaK, 76, 80 Drug target features, 4 E Erythromycin resistance methyltransferase (Erm) enzyme isolation, 227–228 Escherichia coli, 5,12, 38, 68, 76, 98, 120, 134, 189, 191, 217, 245 Essential genes, 4, 7 Ethidium bomide, 20, 177, 188 Ethidium bromide accumulation assay, 191–193 F Fatty acid biosynthesis, 205–206 Flow cytometry, 176–177 Fluorescence polarization assay, 218–219 Fluorescent dyes, 179 Fluorography, 137 Four-part assay for ribosomal subunit formation, 65 Fractional inhibitory concentration method (FIC), 196–197, 201 H Haemophilus influenzae, 2, 54, 68, 189, 191 HeLa cell culture, 93 High throughput screening assay, 45

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274 His-tagged proteins, 14, 30, 41, 124 Human cell extracts, 100 I IC50 , 61, 65 K Kinetoplast DNA, 12 L Large unilamellar vesicle (LUV) assays, 162–163 Lipid quantitation assay, 162 Lipopolysaccharides (LPS), 143–144 Liquid scintillation counting, 32, 67, 77, 79, 147, 149, 199, 211 LpxC deacetylase assay, 147 M MALDI mass spectrophometry, 229 Membrane permeability and potential measurements, 180–181 Membrane preparation, 135, 136 Messenger RNA (mRNA), 88 Miicroorganisms, Bacillus subtilis , 26 Escherichia coli, 5,12, 38, 68, 76, 98, 120, 134, 189, 191, 217, 245 Haemophilus influenzae, 2, 54, 68, 189, 191 Pseudomonas aeruginosa, 6, 188 Staphylococcus aureus, 38, 54, 66, 68, 135 Streptococcus pneumoniae, 19, 54, 68 Minimal inhibitory concentration (MIC), 57, 170, 194–196 Minimal media, 55–56, 64, 77, 93, 120 Mouse infection, 58 Messenger RNA (mRNA) transcription, 96–97 mtFabH assay, 210–211 O ONPG leakage assays, 164 P Penicillin binding proteins (PBP), 131–132 PBP labeling, 134, 136 PBP assays, 137–139 Peptide deformylase (PDF), 117–118 PDF enzyme assays, PDF coupling enzymes, 122 AAP coupled, 125–127 DPPI coupled, 127–128 FDH coupled, 124–125

Index PDF enzyme preparation, 123–125 Plasmid DNA, 12 Post-antibiotic effect, 67 Primer extension assay, 28, 33 Protonophore assay, 197–199 Pseudomonas aeruginosa, 6, 188 Pulse and chase labeling, 67 Q Quinolones, 12 R Ribosomes, 63, 76, 87, 109, 223 Ribosomal subunit, 64 RNA methyltransferases, 224–225 RNA polymerase, 37–38 RNA polymerase assay, 39, 40 RNA polymerase purification, 42–43 RNA transcription, 96, 228 S S100 supernatant preparation, 57,110 Scintillation proximity assay, 39, 109 SDS polyacrylamide gel electrophoresis (PAGE), 40, 47, 133, 137 Sigma factor purification, 43–44 SPARK assay, 112–114 SPARK assay principle, 108 Staphylococcus aureus, 38, 54, 66, 68, 135 Streptococcus pneumoniae, 19, 54, 68 Sucrose density gradient centrifugation, 67, 77 Synthetic templates, 27, 32 T Thin-layer chromatography (TLC), 147 Time kill kinetics assays, 58, 200 Transfer RNA synthetases, 53–54 Transfer RNA synthetase assay, 57 Tryptophan fluorescence assays, 166–167 Two-component signal transduction (TCST), 215–216 U Ultrafiltration binding assay, 149 Uridine labeling of RNA, 65, 77 Y Yeast extract preparation, 99 Yeast growth media, 93