Enzymology at the Membrane Interface: Intramembrane Proteases [1st Edition] 9780128122143, 9780128122136

Table of contents :
Content:
Series PagePage ii
CopyrightPage iv
ContributorsPages xi-xiii
PrefacePages xv-xixMichael H. Gelb
Chapter One - Biochemical Characterization of Function and Structure of RseP, an Escherichia coli S2P ProteasePages 1-33Y. Hizukuri, K. Akiyama, Y. Akiyama
Chapter Two - Signal Peptidase Enzymology and Substrate Specificity ProfilingPages 35-57R.E. Dalbey, D. Pei, Ö.D. Ekici
Chapter Three - Screening and Characterization Strategies for Nanobodies Targeting Membrane ProteinsPages 59-97S. Veugelen, M. Dewilde, B. De Strooper, L. Chávez-Gutiérrez
Chapter Four - Probing the Activity of Eukaryotic Rhomboid Proteases In VitroPages 99-126B. Cordier, M.K. Lemberg
Chapter Five - Expression, Purification, and Enzymatic Characterization of Intramembrane ProteasesPages 127-155R. Zhou, Y. Shi, G. Yang
Chapter Six - Analyzing Amyloid-β Peptide Modulation Profiles and Binding Sites of γ-Secretase ModulatorsPages 157-183J. Trambauer, A. Fukumori, B. Kretner, H. Steiner
Chapter Seven - Probing the Structure and Function Relationships of Presenilin by Substituted-Cysteine Accessibility MethodPages 185-205T. Tomita
Chapter Eight - A New Method to Determine the Transmembrane Conformation of Substrates in Intramembrane Proteolysis by Deep-UV Resonance Raman SpectroscopyPages 207-228J.W. Cooley, A. Abdine, M. Brown, J. Chavez, B. Lada, R.D. JiJi, I. Ubarretxena-Belandia
Chapter Nine - An Inducible Reconstitution System for the Real-Time Kinetic Analysis of Protease Activity and Inhibition Inside the MembranePages 229-253R.P. Baker, S. Urban
Chapter Ten - Production of Recombinant Rhomboid ProteasesPages 255-278E. Arutyunova, R. Panigrahi, K. Strisovsky, M.J. Lemieux
Chapter Eleven - Mechanism and Inhibition of Rhomboid ProteasesPages 279-293K. Strisovsky
Chapter Twelve - Enzymatic Assays for Studying Intramembrane ProteolysisPages 295-308D.M. Bolduc, D.J. Selkoe, M.S. Wolfe
Chapter Thirteen - Methods for Structural and Functional Analyses of Intramembrane Prenyltransferases in the UbiA SuperfamilyPages 309-347Y. Yang, N. Ke, S. Liu, W. Li
Chapter Fourteen - Functional Study of the Vitamin K Cycle Enzymes in Live CellsPages 349-394J.-K. Tie, D.W. Stafford
Chapter Fifteen - Activity Assays for Rhomboid ProteasesPages 395-437E. Arutyunova, K. Strisovsky, M.J. Lemieux
Author IndexPages 439-464
Subject IndexPages 465-474

Citation preview

METHODS IN ENZYMOLOGY Editors-in-Chief

ANNA MARIE PYLE Departments of Molecular, Cellular and Developmental Biology and Department of Chemistry Investigator, Howard Hughes Medical Institute Yale University

DAVID W. CHRISTIANSON Roy and Diana Vagelos Laboratories Department of Chemistry University of Pennsylvania Philadelphia, PA

Founding Editors

SIDNEY P. COLOWICK and NATHAN O. KAPLAN

Academic Press is an imprint of Elsevier 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States 525 B Street, Suite 1800, San Diego, CA 92101–4495, United States The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom 125 London Wall, London, EC2Y 5AS, United Kingdom First edition 2017 Copyright © 2017 Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. ISBN: 978-0-12-812213-6 ISSN: 0076-6879 For information on all Academic Press publications visit our website at https://www.elsevier.com/

Publisher: Zoe Kruze Acquisition Editor: Zoe Kruze Editorial Project Manager: Helene Kabes Production Project Manager: Magesh Kumar Mahalingam Cover Designer: Greg Harris Typeset by SPi Global, India

CONTRIBUTORS A. Abdine Icahn School of Medicine at Mount Sinai, New York, NY, United States K. Akiyama Institute for Frontier Life and Medical Sciences, Kyoto University, Kyoto, Japan Y. Akiyama Institute for Frontier Life and Medical Sciences, Kyoto University, Kyoto, Japan E. Arutyunova Faculty of Medicine and Dentistry, Membrane Protein Disease Research Group, University of Alberta, Edmonton, AB, Canada R.P. Baker Johns Hopkins University School of Medicine, Baltimore, MD, United States D.M. Bolduc Ann Romney Center for Neurologic Diseases, Brigham and Women’s Hospital, Harvard Medical School, Boston, MA, United States M. Brown University of Missouri, Columbia, MO, United States J. Chavez Icahn School of Medicine at Mount Sinai, New York, NY, United States L. Cha´vez-Guti
errez University of Leuven; VIB Center for Brain and Disease, Leuven, Belgium J.W. Cooley University of Missouri, Columbia, MO, United States B. Cordier Zentrum f€ ur Molekulare Biologie der Universit€at Heidelberg (ZMBH), DKFZ-ZMBH Allianz, Heidelberg, Germany R.E. Dalbey The Ohio State University, Columbus, OH, United States B. De Strooper University of Leuven; VIB Center for Brain and Disease, Leuven, Belgium; UCL Institute of Neurology, London, United Kingdom M. Dewilde University of Leuven; VIB Center for Brain and Disease, Leuven, Belgium € O.D. Ekici The Ohio State University, Newark, OH, United States

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A. Fukumori Biomedical Center (BMC), Metabolic Biochemistry, Ludwig-Maximilians-University Munich; German Center for Neurodegenerative Diseases (DZNE), Munich, Germany Y. Hizukuri Institute for Frontier Life and Medical Sciences, Kyoto University, Kyoto, Japan R.D. JiJi University of Missouri, Columbia, MO, United States N. Ke New England Biolabs, Ipswich, MA, United States B. Kretner Biomedical Center (BMC), Metabolic Biochemistry, Ludwig-Maximilians-University Munich; German Center for Neurodegenerative Diseases (DZNE), Munich, Germany B. Lada University of Missouri, Columbia, MO, United States M.K. Lemberg Zentrum f€ ur Molekulare Biologie der Universit€at Heidelberg (ZMBH), DKFZ-ZMBH Allianz, Heidelberg, Germany M.J. Lemieux Faculty of Medicine and Dentistry, Membrane Protein Disease Research Group, University of Alberta, Edmonton, AB, Canada S. Liu Washington University School of Medicine, St. Louis, MO, United States W. Li Washington University School of Medicine, St. Louis, MO, United States R. Panigrahi Faculty of Medicine and Dentistry, Membrane Protein Disease Research Group, University of Alberta, Edmonton, AB, Canada D. Pei The Ohio State University, Columbus, OH, United States D.J. Selkoe Ann Romney Center for Neurologic Diseases, Brigham and Women’s Hospital, Harvard Medical School, Boston, MA, United States Y. Shi Ministry of Education Key Laboratory of Protein Science, Tsinghua-Peking Joint Center for Life Sciences, Beijing Advanced Innovation Center for Structural Biology, School of Life Sciences, Tsinghua University, Beijing, China D.W. Stafford University of North Carolina at Chapel Hill, Chapel Hill, NC, United States H. Steiner Biomedical Center (BMC), Metabolic Biochemistry, Ludwig-Maximilians-University Munich; German Center for Neurodegenerative Diseases (DZNE), Munich, Germany

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K. Strisovsky Institute of Organic Chemistry and Biochemistry, Academy of Sciences of the Czech Republic, Prague, Czech Republic J.-K. Tie University of North Carolina at Chapel Hill, Chapel Hill, NC, United States T. Tomita Laboratory of Neuropathology and Neuroscience, Graduate School of Pharmaceutical Sciences, The University of Tokyo, Tokyo, Japan J. Trambauer Biomedical Center (BMC), Metabolic Biochemistry, Ludwig-Maximilians-University Munich, Munich, Germany I. Ubarretxena-Belandia Icahn School of Medicine at Mount Sinai, New York, NY, United States S. Urban Johns Hopkins University School of Medicine, Baltimore, MD, United States S. Veugelen University of Leuven; VIB Center for Brain and Disease, Leuven, Belgium M.S. Wolfe Ann Romney Center for Neurologic Diseases, Brigham and Women’s Hospital, Harvard Medical School, Boston, MA, United States G. Yang Ministry of Education Key Laboratory of Protein Science, Tsinghua-Peking Joint Center for Life Sciences, Beijing Advanced Innovation Center for Structural Biology, School of Life Sciences, Tsinghua University, Beijing, China Y. Yang Washington University School of Medicine, St. Louis, MO, United States R. Zhou Ministry of Education Key Laboratory of Protein Science, Tsinghua-Peking Joint Center for Life Sciences, Beijing Advanced Innovation Center for Structural Biology, School of Life Sciences, Tsinghua University, Beijing, China

PREFACE Interfacial enzymology is concerned with enzymes that must access their substrates from a boundary between two phases (an interface), the most common type being the lipid–water interface of biological membranes (Berg & Jain, 2002). Interfacial enzymes are distinct from noninterfacial enzymes (Gelb, Jain, Hanel, & Berg, 1995). The latter includes enzymes in the water layer that act on substrates in the water layer. One example is hexokinase, which is a water-soluble enzyme that acts on the highly water-soluble substrate glucose. Noninterfacial also include enzymes in the membrane that act on watersoluble substrates. One example is the phosphodiesterase in the visual transduction cycle that hydrolyzes cyclic GMP. These noninterfacial enzymes have in common that their active site should access substrate in the water layer even though one of the enzymes is an integral membrane protein. Presumably the active site is well exposed to the water layer even if the enzyme is bound to membranes. Noninterfacial enzymes are sensitive to the concentration of substrate in the aqueous phase (moles of substrate per volume of aqueous phase). Rate equations describing noninterfacial enzymes are composed of rate constants and aqueous phase concentrations of interacting partners (enzyme and substrate). By contrast, interfacial enzymes must access their substrates from the interface, and the rate equations that describe their action depend on rate constants and the moles of substrate per volume of interface. Among interfacial enzymes, we also include those that access their substrates from the membrane core, and in these cases the rate equation contains terms for the concentration of substrate in the membrane core. For enzymes that act on highly water-insoluble substrates, for example, phospholipids with long fatty acyl chains or transmembrane segments of proteins, the concentration of substrate in the aqueous phase is vanishingly small, so much so that the enzyme is forced to access its substrate from the membrane phase. These are the interfacial enzymes. Strictly speaking, it is not possible to conclude that an enzyme is interfacial based on solubility arguments alone. Proof comes from the observation of processive behavior. Consider the enzyme secreted phospholipase A2. This is a water-soluble enzyme that absorbs onto the surface of phospholipid vesicles and thus exists in a water-soluble state (E) and an interfacial state (E*). It has been experimentally demonstrated under some conditions that enzyme prebound to vesicles of one type of phospholipid is not able to act on vesicles of a different xv

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phospholipid added later (under conditions where there is no intervesicle transfer of phospholipids), yet if enzyme is added to a premixture of both vesicles, it acts on both types of phospholipids (Gelb et al., 1995). Enzyme bound to the first vesicle is able to remain bound to the vesicle and catalyze the hydrolysis of several phospholipids before leaving the interface. If the enzyme was acting on the trace amount of phospholipid in the water phase, this type of processive process would clearly not be possible. These results also show that binding of enzyme to the interface (E to E*) is not the same step as loading the active site of the enzyme with substrate (Michaelis complex formation, E* to E*S). If these steps were the same, release of product from the enzyme’s active site would necessarily result in release of enzyme from the membrane interface to the water layer and processive behavior would not be possible. Processive behavior of interfacial enzymes has been called “scooting” by Mahendra Jain and coworkers, whereas nonprocessive behavior is called “hopping” (Jain & Berg, 1989). An interesting dilemma results with an enzyme that acts on a substrate that has significant solubility in both the membrane and aqueous phases. One example is platelet-activating factor acetylhydrolase that cleaves the ester of a phospholipid containing a short acetyl group instead of a longchain fatty acyl group. The enzyme is water soluble but found in vivo bound tightly to lipoproteins in blood. Does the enzyme access its substrate only in the aqueous phase (for example, the membrane-bound enzyme has its active site exposed mainly to the water layer) or does the enzyme access its substrate only in the lipoprotein (for example, its active site is not well exposed to the aqueous phase)? The question is nontrivial to answer because the substrate readily partitions between the aqueous and membrane phases. Thus, at equilibrium, the concentration of substrate in the membrane phase is equal to the concentration of substrate in the aqueous multiplied by the partition equilibrium constant. Also during equilibrium, variation of the concentration of substrate in one phase leads to a proportional variation of substrate in the other phase. Any steady-state rate equation that is written in terms of moles of substrate in the aqueous phase divided by the volume of the aqueous phase (conventional concentration) can be rewritten in terms of the moles of substrate in the membrane phase divided by the volume of membrane phase (interfacial concentration) times a constant. The two equations are mathematically equivalent, and it is thus impossible to design any steady-state kinetic experiment to determine which equation applies, i.e., whether the enzyme is interfacial or not. The problem has been solved for platelet-activating factor acetylhydrolase by studying the kinetics during

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the presteady-state phase in which the substrate has not yet equilibrated between membrane and aqueous phases (Min et al., 1999). This study showed that the enzyme operates on platelet-activating factor in the aqueous phase and is thus not an interfacial enzyme. One may wonder whether a membrane-bound signaling receptor that is gated by a ligand that can exist in the membrane and aqueous phases interacts with ligand in the membrane phase (interfacial receptor) or the aqueous phase (noninterfacial receptor). This question has never been answered. For interfacial enzymes that display fast turnover numbers, the catalytic cycle may be limited by the rate of exchange of substrate between substrate aggregates containing bound enzyme. For example, some secreted phospholipases A2 display a turnover number of
100 s1. If the enzyme is bound to a small unilamellar vesicle containing say 500 phospholipids, substrate would become exhausted in just a few seconds. To continue, enzyme must move to a new vesicle or there must be intervesicle exchange of phospholipids, and these processes may become rate-limiting for steady-state turnover. In this case the steady-state parameters do not reflect the true catalytic properties of the enzyme but rather those of substrate and enzyme intervesicle dynamics. This problem is especially pertinent to phospholipid–detergent mixed micelles that contain only a few phospholipids per particle (Dennis, Cao, Hsu, Magrioti, & Kokotos, 2011). For enzymes such as secreted phospholipase A2, there are two components to their substrate specificity. The first issue is what are the structure–function relationships that determine the affinity of the enzyme for the membrane surface (E to E*). Once bound to the interface, then comes the issue of the structural requirements of the interfacial substrate for binding to the catalytic site of E* to give the Michaelis complex E*S. For example, group IIA secreted phospholipase A2 does not appreciably bind to vesicles that lack anionic phospholipids (for example, vesicles rich in phosphatidylcholine), yet once bound to anionic phospholipid vesicles, phosphatidylcholine that may be present in the same vesicles are good substrates for the enzyme (thus, E* + S Ð E*S is favorable) (Gelb et al., 1995). Often substrate specificity studies of interfacial enzymes are carried out in ways that do not allow the deconvolution of these two processes and are thus almost impossible to interpret and to extrapolate from in vitro to in vivo settings. Inhibition of interfacial enzymes is more difficult to study than with noninterfacial enzymes (Gelb et al., 1995). It is possible for inhibitors to act through nonspecific mechanisms, for example, compounds that partition into the membrane interface and change the physical properties of the

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interface in ways that promote enzyme desorption to the aqueous phase (E* to E). More interesting and useful inhibitors are those that bind specifically to the catalytic site (or other site) on the enzyme (E* + I gives E*I); these are analogous to inhibitors of noninterfacial enzymes. Early studies of phospholipase A2 inhibitors were plagued by the common occurrence of nonspecific, membrane-perturbing inhibitors. For example, annexins were initially characterized as phospholipase A2 inhibitors, but later studies showed that annexins form a tightly packed array on the membrane surface and simply occlude the interfacial enzyme from binding to the vesicle to access its substrate. Thus, annexins, at least in vitro, are inhibitors of virtually all interfacial enzymes that must undergo an E to E* transition as part of its catalytic cycle. Volume 583 of Methods in Enzymology is focused on interfacial enzymes that act on lipid substrates. Methods for detecting and quantifying the binding of proteins and enzymes to membranes in vitro are covered in Chapters 1, 9, 10, and 11. Methods for detection of interfacial binding of proteins and enzymes to membranes in living cells are covered in Chapters 2, 4, 6, 13, and 15. Some interfacial enzymes form protein–protein complexes, and this is described in Chapters 8 and 14. Studies to evaluate the substrate specificity of interfacial enzymes on natural members are covered in Chapter 5. Novel spectroscopic studies for providing structural insight into protein–membrane binding are provided in Chapters 7, 9, 11, and 12. Methods for production of recombinant interfacial enzymes are given in Chapters 3 and 6. Volume 584 of Methods in Enzymology is focused on proteolytic enzymes that access their transmembrane peptide substrates in the membrane phase (interfacial proteases). Biochemical characterization of bacterial and eukaryotic transmembrane proteases is covered in Chapters 1, 2, 4, 5, 6, 7, 10, and 15. Kinetic studies including substrate specificity and inhibition are included in Chapters 2, 4, 5, 6, 9, 11, 12, and 15. Studies that probe the structure of transmembrane proteases are given in Chapters 3, 6, 8, and 13. Production of recombinant transmembrane proteases is the specific focus of Chapters 5 and 10. In summary, interfacial enzymes are an important subset of enzymes. Early studies were focused on membranes that act at the lipid–water interface. More recently a new class of proteases that act on transmembrane protein segments have been discovered in several organisms. Special methods are required for the characterization of interfacial enzymes including their basic features of substrate specificity and inhibition. Structural studies are challenging because the enzyme acts in an environment that is not amenable to conventional techniques for determining molecular structure. MICHAEL H. GELB University of Washington, Seattle, WA, United States

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REFERENCES Berg, O. G., & Jain, M. K. (2002). Interfacial enzyme kinetics. West Sussex, England: Wiley. Dennis, E. A., Cao, J., Hsu, Y.-H., Magrioti, V., & Kokotos, G. (2011). Phospholipase A2 enzymes: Physical structure, biological function, disease implication, chemical inhibition, and therapeutic intervention. Chemical Reviews, 111, 6130–6185. http://dx.doi. org/10.1021/cr200085w. Gelb, M. H., Jain, M. K., Hanel, A. M., & Berg, O. (1995). The interfacial enzymology of glycerolipid lipases: Lessons from secreted phospholipases A2. Annual Review of Biochemistry, 64, 653–688. Jain, M. K., & Berg, O. (1989). The kinetics of interfacial catalysis by phospholipase A2 and regulation of interfacial activation: Hopping versus scooting. Biochimica et Biophysica Acta, 1002, 127–156. Min, J. H., Jain, M. J., Wilder, C., Paul, L., Apitz-Castro, R., Aspleaf, D. C., et al. (1999). Membrane-bound plasma platelet activating factor acetylhydrolase acts on substrate in the aqueous phase. Biochemistry, 38, 12935–12942.

CHAPTER ONE

Biochemical Characterization of Function and Structure of RseP, an Escherichia coli S2P Protease Y. Hizukuri, K. Akiyama, Y. Akiyama1 Institute for Frontier Life and Medical Sciences, Kyoto University, Kyoto, Japan 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. The Use of MBP-Tagged Model Substrates for Analysis of Proteolytic Activity of RseP 2.1 The In Vivo Cleavage Assay 2.2 Cleavage of the Model Substrates by Purified RseP In Vitro 3. In Vitro Analysis of Proteolysis of SPs by RseP 3.1 Handling of Small Membrane Proteins Like an SP 3.2 Cleavage of Synthetic SPs With Small Tags In Vitro 3.3 Proteolysis of Authentic SPs Inside the Membrane 4. Structural Analysis of RseP Using Thiol-Specific Modification Reagents 4.1 Evaluation of the Environment Around the Protease Active-Site Region of RseP 4.2 Evaluation of Solvent Accessibility of the Periplasmic Domain of RseP 4.3 Determination of In Vivo Substrate Cleavage Sites Using mal-PEG Modification 5. In Vivo Cross-Linking Analysis to Probe the Interaction of RseP With Its Substrate in the Membrane 5.1 In Vivo Photo Cross-Linking 5.2 Disulfide Cross-Linking Acknowledgments References

2 4 5 8 11 11 13 14 18 18 22 25 27 27 29 32 32

Abstract Intramembrane-cleaving proteases (I-CLiPs) are a group of membrane-associated proteases with a unique feature: they are believed to cleave their substrate within the hydrophobic lipid bilayer, even though peptide bond hydrolysis requires a water molecule. Escherichia coli RseP, which belongs to the S2P zinc metalloprotease family of I-CLiPs, plays an essential role in activation of a cell envelope stress response through cleavage of anti-σE protein RseA, a single-span transmembrane protein. A recent study

Methods in Enzymology, Volume 584 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2016.09.044

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2017 Elsevier Inc. All rights reserved.

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showed that it also cleaves remnant signal peptides generated upon membrane translocation of secretory proteins. Here, we describe several methods for characterization of the proteolytic functions and structure of RseP mainly in vivo, including a proteolytic activity assay using model substrates, an in vitro analysis of cleavage of signal peptides in a detergent solution and in the membrane vesicles, structural analysis of membraneembedded RseP based on the thiol modifiability of introduced cysteine residues, and the protein interaction analysis by in vivo cross-linking protocols.

1. INTRODUCTION In living cells, many proteases function at membranes. Most membrane-associated proteases have their catalytic site exposed to an aqueous compartment and act on substrates in a manner similar to that of watersoluble proteases. Contrary to this “classical” type of membrane proteases, a new type of membrane-associated proteases was discovered approximately two decades ago: their conserved catalytic sites are apparently membrane embedded and likely execute proteolysis inside the lipid bilayer (Rawson et al., 1997). This novel class of proteases was named intramembranecleaving proteases (I-CLiPs) (Weihofen & Martoglio, 2003; Wolfe, 2009), and they have been found to perform important functions in a variety of cellular events through the process called regulated intramembrane proteolysis (RIP) both in prokaryotic and eukaryotic cells (Brown, Ye, Rawson, & Goldstein, 2000; Urban, 2013). I-CLiPs are categorized into three classes: presenilin (the catalytic subunit of γ-secretase)/signal peptide peptidase (aspartyl protease), rhomboid protease (serine protease), and S2P protease (metalloprotease) based on their catalytic mechanism and amino acid sequence homology. The three-dimensional structures of proteases belonging to each of these three classes have been reported (Bai et al., 2015; Feng et al., 2007; Sun, Li, & Shi, 2016; Wang, Zhang, & Ha, 2006). RseP, formerly known as YaeL, is an S2P family zinc metalloprotease ortholog in Escherichia coli and is involved in activation of the σE-pathway of the extracytoplasmic stress response, which plays important roles in bacterial survival when bacterial cells are exposed to a variety of cell surface stressors that cause damage to the outer membrane (Ades, 2008; Chen & Zhang, 2010; Kanehara, Akiyama, & Ito, 2001; Kroos & Akiyama, 2013) (Fig. 1A). Under unstressful conditions, σE, an extracytoplasmic function σ factor, is trapped by the N-terminal cytoplasmic region of anti-σ factor

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Fig. 1 Functions of a bacterial S2P family protease, RseP. (A) RseP activates the σE pathway of extracytoplasmic stress response by cleaving anti-σE protein RseA. Some kinds of envelope stressors promote sequential cleavage of RseA by DegS (site-1 cleavage) and subsequently by RseP (site-2 cleavage). (B) RseP also acts to proteolytically remove remnant signal peptides (SPs) left over in the cytoplasmic membrane. SPs generated by Leader peptidase (Lep)-mediated processing of presecretory proteins are cleaved by RseP.

RseA, a type II (Nin–Cout) single-span transmembrane protein, and is kept inactivated. The stress cues cause accumulation of malfolded outermembrane proteins and derivatives of lipopolysaccharides in the periplasmic space, and this process induces cleavage of RseA in its periplasmic region by

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the membrane-tethered serine protease DegS (site-1 cleavage) (Lima, Guo, Chaba, Gross, & Sauer, 2013). This first cleavage triggers the RseP-catalyzed second cleavage of the transmembrane region of RseA (site-2 cleavage) and liberation of the RseA cytoplasmic domain complexed with σE from the membrane. The cytoplasmic domain of RseA is degraded in the cytoplasm by proteases such as ClpXP, leading to eventual activation of σE. RseP cannot act on intact RseA and cleaves it only after DegS truncates the periplasmic part of RseA (Alba, Leeds, Onufryk, Lu, & Gross, 2002; Kanehara, Ito, & Akiyama, 2002). The site-1 cleavage dependence of the site-2 cleavage ensures stress-induced activation of σE. This regulated proteolysis is thought to be controlled by tandemly arranged two PDZ domains (PDZ tandem) in the periplasmic region of RseP (Hizukuri & Akiyama, 2012; Inaba et al., 2008; Kanehara, Ito, & Akiyama, 2003), in a way that the PDZ tandem serves as a size exclusion filter to prevent intact RseA (which has a bulky periplasmic region) from accessing the recessed catalytic site in the membrane domain of RseP (Hizukuri et al., 2014). This size-dependent substrate discrimination mechanism would also operate to cleave remnant signal peptides (SPs) only after they are detached from the mature region of secretory proteins (Saito et al., 2011) (Fig. 1B). In addition, a recent study showed that a β-hairpin-like structure (MRE β-loop) located close to the active site in the RseP membrane domain is implicated in substrate cleavage by specifically recognizing the transmembrane region of substrates (Akiyama et al., 2015). In this chapter, we describe several approaches that we have used to analyze RseP. Biochemical analyses that are carried out in detergent extracts as well as structural research such as X-ray crystallography undoubtedly have provided important information, but we believe that functional and structural characterization of an enzyme under more physiological conditions, i.e., in the membrane, is necessary to understand its physiological roles. The proposed approaches should help to study other I-CLiPs and membrane-embedded enzymes.

2. THE USE OF MBP-TAGGED MODEL SUBSTRATES FOR ANALYSIS OF PROTEOLYTIC ACTIVITY OF RseP To conduct a quantitative or kinetic analysis of the proteolytic activity of an I-CLiP in vivo, it is desirable to quantitatively detect the proteolytic products of a substrate protein. In the case of RseP, however, the cytoplasmic fragment of RseA generated by RseP is unstable and rapidly degraded by other cellular proteases such as ClpXP. In addition, because intact RseA is

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first cleaved by DegS on the periplasmic side, and this first cleavage is usually a prerequisite for the following intramembrane cleavage by RseP, formation of the RseP-cleaved product from intact RseA depends on these two proteases. These features of the in vivo RseA degradation make quantitative analysis of the RseP-catalyzed cleavage of RseA difficult. Thus, we constructed a model substrate that allows for quantitative analysis of the RseP activity in vivo (Akiyama, Kanehara, & Ito, 2004) (Fig. 2A). In this substrate (HA-MBP-RseA140), the cytoplasmic region of RseA was replaced with a stably folding maltose-binding protein (MBP; M.W.
41,000) domain with an N-terminal hemagglutinin (HA) tag. The MBP domain stabilizes the RseA-cleaved product in the cytoplasm, and the HA tag facilitates its immunological detection. In addition, the periplasmic region was truncated to mimic DegS-catalyzed site-1 cleavage. We also designed a model substrate with a similar construct (HA-MBP-RseA(LY1)140) in which the transmembrane region of RseA in HA-MBP-RseA140 was replaced by the first transmembrane segment (LY1) of lactose permease (LacY). Although LY1 is not a physiological substrate of RseP, we found that this model substrate is efficiently cleaved by RseP and is more suitable for quantitative analysis of the RseP activity in vivo (see later). The use of a stable tag domain such as MBP should be useful for analysis of some other I-CLiPs, especially when the cleavage product of their substrate is unstable. This approach may also allow for easier separation and detection of cleaved products in a standard SDS-PAGE system and for easier purification of substrate membrane proteins (see Section 2.2).

2.1 The In Vivo Cleavage Assay For in vivo cleavage assays to evaluate the proteolytic activity of RseP against the HA-MBP-fused model substrate in living cells, we usually use the E. coli rseP rseA double-disrupted strains carrying two compatible plasmids, one expressing RseP-His6-Myc or its variant and the other expressing the model substrate such as HA-MBP-RseA140, HA-MBP-RseA(LY1)140, or HA-MBP-SPBla-Bla (see later). RseP-His6-Myc has a bipartite His6-Myc tag at its C-terminus; the hexahistidine tag (His6) enables purification on a Ni-NTA column, and the Myc tag is used for detection by immunoblotting or immunoprecipitation with an anti-Myc antibody. As expected, HA-MBP-RseA140 was cleaved by RseP in a DegSindependent manner in vivo, and the cytoplasmic cleavage product (CL) was detected (Fig. 2, lanes 1–4) (Akiyama et al., 2004). Nevertheless, the CL fragment of this substrate was still subject to partial degradation because

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C 140 RseA TM

C 216

DegS(1st)

C

C

B

HA-MBP-SPBla-Bla

HA-MBP-RseA140 1

2

3

4

5

6

7

8

(kDa) 1

UC CL

αHA

81.8

Full

68.4

αMBP rseP degS

2

+ +

Δ +

+ Δ

Δ Δ

RseATM

+ +

Δ +

+ Δ

Δ Δ

55.0 UC CL

rseP

+

Δ

41.6

αHA

LacYTM1

Fig. 2 RseP-dependent cleavage of the MBP-fused model substrates in vivo. (A) Schematic diagrams of various model substrates. (B) Immunological detection of the cleaved N-terminal products of the model substrates. A plasmid encoding HA-MBP-RseA140 (left half ) or HA-MBP-RseA(LY1)140 (right half ) was introduced into a ΔrseA strain (lanes 1 and 5) as well as into its derivative with an additional deletion of rseP (lanes 2 and 6), degS (lanes 3 and 7), or both (lanes 4 and 8). Protein samples were analyzed by standard SDS-PAGE and anti-HA or anti-MBP immunoblotting. RseP-uncleaved (UC) and RseP-cleaved (CL) forms of the substrates are indicated. (C) Cleavage of HA-MBP-SPBla-Bla. Proteins were analyzed as in (A). Full, an intact substrate before processing by Lep. Modified from Akiyama, Y., Kanehara, K., & Ito, K. (2004). RseP (YaeL), an Escherichia coli RIP protease, cleaves transmembrane sequences. EMBO Journal, 23(22), 4434–4442; Saito, A., Hizukuri, Y., Matsuo, E., Chiba, S., Mori, H., Nishimura, O., et al. (2011). Post-liberation cleavage of signal peptides is catalyzed by the site-2 protease (S2P) in bacteria. Proceedings of the National Academy of Sciences of the United States of America, 108(33), 13740–13745.

Functional and Structural Analysis of RseP

7

ClpXP still recognizes a newly generated C-terminus of this fragment and degrades it according to its stabilization in a clpP-disrupted strain (Fig. 2) (Akiyama et al., 2015). HA-MBP-RseA(LY1)140 (Akiyama et al., 2004), another model substrate, also underwent efficient RseP cleavage and accumulated its N-terminal fragment more stably (Fig. 2B, lanes 5 and 7). This model substrate allows for evaluation of a proteolytic activity of RseP (and mutants) quantitatively even in a clpP+ host strain. We found that RseP can cleave, in addition to RseA, various SPs generated from precursor forms of secretory proteins (Saito et al., 2011). Accompanying translocation of the presecretory protein across the membrane, it is processed by Leader peptidase (Lep) to generate a remnant SP that is left in the membrane as a type II integral membrane protein. Because the SPs are small peptides (20 amino acid residues), it is generally difficult to detect them, especially when they are generated in vivo. We found that attachment of the HA-MBP protein tag to the N-terminus of secretory protein precursors enables detection and analysis of SPs. For example, HA-MBP-SPBla-Bla is a precursor form of β-lactamase (Bla) with an HA-MBP domain attached to the N-terminus of its SP (Fig. 2A). When this model protein was expressed, the SP with the N-terminal HA-MBP tag (HA-MBP-SPBla-Bla) was generated by the action of Lep (UC in Fig. 2C) and underwent the RseP-dependent cleavage (CL in Fig. 2C, lane 1). Using essentially the same approach, we also demonstrated that RasP, an RseP ortholog in Bacillus subtilis, can cleave SPs of B. subtilis secretory proteins (Saito et al., 2011). A typical protocol for the in vivo cleavage assay is as follows: 1. Grow cells at 30°C until the early- or mid-log phase. In the case of RseP, the use of a minimal medium (such as the M9 medium) could yield better results as compared with a rich medium. 2. Chill the cell cultures on ice and precipitate total cellular protein by mixing equal volumes of the culture and ice-cold 10% trichloroacetic acid (TCA). After incubation of the samples at 0°C for 20 min, protein pellets are collected by centrifugation (20,000  g, 5 min at 4°C) and washed with cold acetone. 3. Resuspend the precipitated proteins in 1  SDS sample buffer (containing 10% 2-mercaptoethanol (ME)) by mixing vigorously at room temperature for 30 min followed by incubation at 37°C for 5 min. 4. Analyze the samples by SDS-PAGE (usually, in a Laemmli gel) and by immunoblotting (fusion of the MBP domain to a target protein causes 41-kDa upshift in an apparent size on an SDS-polyacrylamide gel).

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Antibody-bound proteins are visualized on a lumino image analyzer using a general chemiluminescent method.

2.2 Cleavage of the Model Substrates by Purified RseP In Vitro In addition to in vivo assays, in vitro analysis is crucial for elucidation of the detailed mechanism of substrate proteolysis by RseP. Here, we describe a simple assay of proteolytic cleavage of the MBP-fused RseA model substrate by RseP in a purified system. In the study of the rhomboid protease (Dickey, Baker, Cho, & Urban, 2013; Moin & Urban, 2012) and γ-secretase (presenilin) (Bolduc, Montagna, Gu, Selkoe, & Wolfe, 2016), elegant reconstitution assays have been devised, which allow for real-time kinetic analysis of substrate proteolysis within the lipid bilayer (Langosch, Scharnagl, Steiner, & Lemberg, 2015). Unfortunately, a similar assay system is not yet available for an S2P family protease and should be established in the future. 2.2.1 Purification of RseP and of the RseA-Derived Model Substrates For purification of RseP and of the model substrates, RseP is overproduced using a strain carrying a plasmid (a derivative of a pUC-based vector) expressing the wild-type (WT) RseP-His6-Myc protein or its variants. The RseA-derived model substrates (e.g., His6-MBP-RseA140) are overproduced using an rseP and rseA double-deletion strain carrying a plasmid (a derivative of a pACYC184-based vector) expressing a substrate; the rseP-disrupted strain is used as a host to prevent cleavage of substrates by endogenous RseP during cultivation. The N- or C-terminally attached His6 tag enables one-step purification by Ni-NTA affinity chromatography. Although His6-tagged RseA140 proved intractable due to its high tendency for aggregation, replacement of the RseA cytoplasmic domain with an MBP protein allowed us to purify a sufficient amount of the substrate protein. The MBP tag may be a good choice for easier purification of some other membrane proteins. A Coomassie Brilliant Blue (CBB) R-250-stained gel after SDS-PAGE with purified RseP-His6-Myc, its H22F active-site mutant (His22 is a catalytic histidine coordinating a zinc ion), and His6-MBP-RseA140 is shown in Fig. 3A (Akiyama et al., 2004). Typical protocols for the purification of RseP and of the MBP-fused model substrates are as follows: 1. Grow cells overexpressing a target protein at 30°C in 1–3 L of the L medium until the mid-log phase.

9

Functional and Structural Analysis of RseP

B

A 3

200 131

RseP UC

78

CL

1

2

3

4

5

4

8

6

7

8

9

0

8

0

8

CBB

2

αMBP

1

UC 39

CL Time (h)

31

RseP

0

1

2 WT

WT+PT

H22F

His6-MBP-RseA140

WT

RseP-His6-Myc

H22F

CBB

His6-MBP-RseA140

Fig. 3 In vitro proteolysis of the RseA model substrate catalyzed by RseP. (A) Purified preparations of RseP-His6-Myc and the RseA model substrate proteins. A 0.35-mg aliquot of each purified sample of RseP-His6-Myc (lane 1), RseP(H22F)-His6-Myc (lane 2), and His6MBP-RseA140 (lane 3) was separated by SDS-PAGE in a 10% Laemmli gel with CBB R-250 staining. (B) Cleavage of His6-MBP-RseA140 by RseP. His6-MBP-RseA140 was incubated with RseP-His6-Myc (lanes 1–7) or RseP(H22F)-His6-Myc (lanes 8 and 9). Fivemillimolar 1,10-phenanthroline (PT) was added to inhibit the RseP activity (lanes 6 and 7). Upper panel: CBB staining, lower panel: anti-MBP immunoblotting. Modified from Akiyama, Y., Kanehara, K., & Ito, K. (2004). RseP (YaeL), an Escherichia coli RIP protease, cleaves transmembrane sequences. EMBO Journal, 23(22), 4434–4442.

2. Harvest the cells by centrifugation, wash with 100 mL of 10 mM Tris–HCl (pH 8.1), centrifuge again, and resuspend in 25 mL of 10 mM Tris–HCl (pH 8.1) with 1 mM phenylmethylsulfonyl fluoride (PMSF, a serine and cysteine protease inhibitor; Nacalai Tesque, Inc., Kyoto, Japan, catalog #: 27327) at 4°C. For purification of a metalloprotease such as RseP, the use of EDTA or other metal chelators should be avoided throughout all steps because it inhibits proteolytic activity. In addition, metal-chelators may hamper the subsequent Ni-NTA affinity purification. 3. Disrupt cells using a French press (8000 psi  2 ), remove cell debris by low-speed centrifugation (1600  g, 10 min at 4°C) and recover the total membrane fraction by ultracentrifugation (100,000  g, 60 min at 4°C).

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4. Resuspend the membranes in 2 mL of a buffer consisting of 10 mM Tris–HCl (pH 8.1), 16% glycerol, and 1 mM PMSF, freeze them in liquid nitrogen, and store in a freezer at –80°C. 5. Thaw the membranes on ice, mix with solubilization buffer (10 mM Tris–HCl (pH 8.1), 10% glycerol, 300 mM KCl, 1% n-dodecyl-β-Dmaltoside (DDM), and 1 mM PMSF), and solubilize by incubation for 1 h on ice with occasional mixing. 6. Remove the insoluble materials by ultracentrifugation (100,000  g, 60 min at 4°C) and load the supernatants onto a Ni-NTA agarose column preequilibrated with wash buffer (10 mM Tris–HCl (pH 8.1), 300 mM KCl, 10% glycerol, and 0.02% DDM). 7. Wash the columns extensively with wash buffer supplemented with 20 mM imidazole and elute the bound proteins using wash buffer with a linear 20–500 mM imidazole gradient. 8. Combine the protein peak fractions, dialyze them extensively against dialysis buffer (10 mM Tris–HCl (pH 8.1), 300 mM KCl, 5% glycerol, and 0.02% DDM) at 4°C, and then remove insoluble materials by ultracentrifugation (100,000  g, 30 min at 4°C). 9. Divide the purified proteins into aliquots, freeze them in liquid nitrogen, and store at –80°C. 2.2.2 The Proteolytic Reaction Typical results of an in vitro proteolytic reaction using the purified WT or H22F mutant form of RseP-His6-Myc and the purified His6-MBP-RseA140 are shown in Fig. 3B (Akiyama et al., 2004). The amount of the MBPcontaining cleaved fragment (CL band) gradually increased during incubation at 37°C in an RseP-dependent manner, while the active-site mutant, H22F, generated no cleaved fragment, showing that the substrate cleavage was catalyzed by RseP. Inclusion of 5 mM zinc chelator 1,10-phenanthroline (PT) almost completely inhibited the activity of RseP. A typical protocol for the in vitro proteolytic reaction is as follows: 1. Mix a purified enzyme (final conc. 20–30 μg/mL) and a substrate protein (15 μg/mL) in 1  RseP buffer (see later) containing 1 mM PMSF. If necessary, add PT (Nacalai Tesque, Inc., catalog #: 26707; final conc. 5 mM) to inhibit RseP activity. Beforehand, 10 RseP buffer (0.5 M Tris–HCl (pH 8.1), 0.2% DDM, 25% glycerol, 50 μM zinc acetate, 100 mM ME) can be prepared and stored at –20°C. 0.1 M zinc acetate in 50 mM Tris–acetate (pH 8.0) is used as a stock solution to prepare 10 RseP buffer.

Functional and Structural Analysis of RseP

11

2. Incubate the reaction mixture at 37°C. Withdraw an aliquot at intervals and mix it with an equal volume of 2  SDS sample buffer containing ME. 3. Proteins are separated by SDS-PAGE (usually in a Laemmli gel) and detected by CBB R-250 staining or immunoblotting. It is also possible to assess the substrate cleavage by mixing the solubilized membrane fraction containing a substrate and purified RseP (Hizukuri & Akiyama, 2012).

3. IN VITRO ANALYSIS OF PROTEOLYSIS OF SPs BY RseP Addition of an MBP protein tag to the N-terminus of an SP allows for detection of an SP and for analysis of its cleavage by RseP in vivo (Section 2.1), but the large tag (50 kDa) as compared with the SP itself (2 kDa) may alter the susceptibility of the SP to cleavage by RseP. We thus conducted an analysis of RseP-catalyzed cleavage using a synthetic SP with a small tag(s) or an authentic SP generated upon in vitro translocation of a model secretory protein into inverted membrane vesicles (IMVs). In this section, we describe procedures of these in vitro assays.

3.1 Handling of Small Membrane Proteins Like an SP First, we will give some tips on detection of small proteins (less than 10 kDa) including SPs by SDS-PAGE. Synthetic SP peptides are often hard to dissolve in aqueous solutions due to their relatively high hydrophobicity. Addition of hydrophilic sequences (e.g., a Myc or Flag tag) at the N- and/or C- terminus is one of the useful ways to facilitate handling of synthetic SPs because not only it increases the molecular mass and solubility of the SP peptides, but also the attached sequences can serve tags for immunological detection. For example, we have added a Myc tag to the N-terminus and a Flag tag to the C-terminus of a synthetic β-lactamase (a periplasmic enzyme) SP (Saito et al., 2011) (Fig. 4A). Indeed, the synthetic Myc-SPBla-Flag peptide was cleaved by purified RseP in vitro. SPBla-Flag, which has only the C-terminal Flag tag, also underwent cleavage by RseP but with lower efficiency as compared with Myc-SPBla-Flag. This difference in cleavage efficiency may be ascribable to lower solubility of SPBla-Flag. The following are some notes on gel electrophoresis and gel-staining procedures for better detection of low-molecular-weight proteins or peptides.

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Y. Hizukuri et al.

Myc-SPBla-Flag

A

RseP N22 (F1)

C21 (F2)

EQKLISEEDL MSIQHFRVALIP12FFAAFCLPVFA HPDYKDDDDK SPBla

Myc

Flag

B 1 2 3 4

5

6

7

8

9 10

RseP-His6-Myc

11 12 (kDa) 250 150 100 75 50 37 25 20 15

Bla

Myc-SP

-Flag

10

F2 F1

5 2

Time (h)

CBB

0

3 12 24

0 12

0 12

0 12

0 12

WT

H22F

WT +PT

–

Fig. 4 Cleavage of a synthetic Bla SP by purified RseP. (A) Amino acid sequence of Myc-SPBla-Flag. N22(F1) and C21(F2) indicate cleavage products of the substrate. (B) The in vitro reaction. Cleavage of the Myc-SPBla-Flag peptide by RseP generated the F1 and F2 fragments. The H22F active-site mutation in RseP and the metal-chelator 1,10-phenanthroline (PT) prevented the formation of cleavage products. The samples were analyzed by means of the 12% Bis-Tris/MES PAGE system (Thermo Fisher Scientific Inc. (Invitrogen (Novex)), Waltham, MA; Nu-PAGE) and CBB G-250 staining. Modified from Saito, A., Hizukuri, Y., Matsuo, E., Chiba, S., Mori, H., Nishimura, O., et al. (2011). Postliberation cleavage of signal peptides is catalyzed by the site-2 protease (S2P) in bacteria. Proceedings of the National Academy of Sciences of the United States of America, 108(33), 13740–13745.

(1) Standard Tris/glycine SDS-PAGE (in a Laemmli gel) tends to cause poor separation of proteins of low-molecular weight even at high acrylamide concentrations. The following gel systems could give better results.

Functional and Structural Analysis of RseP

13

(i) The tricine SDS-PAGE system: e.g., a 10–20% gradient tricine gel or 16% tricine gel with tricine SDS running buffer and tricine SDS sample buffer containing 50 mM dithiothreitol (DTT) (Thermo Fisher Scientific Inc. (Invitrogen (Novex))). Heat samples at 85°C for 2 min before electrophoresis. (ii) The Bis-Tris SDS-PAGE system: e.g., Nu-PAGE 12% Bis-Tris gel with MES SDS running buffer and LDS sample buffer with 50 mM DTT (Thermo Fisher Scientific Inc. (Invitrogen (Novex))). Heat samples at 75°C for 5 min before electrophoresis. (2) It is better to perform electrophoresis at low constant voltage (e.g., 100 V). The use of higher voltage may cause diffused electrophoresis of small proteins. It is also recommended to perform electrophoresis at a low temperature (i.e., in a cold room) because electrophoresis of small proteins may be problematic at elevated temperatures. (3) CBB staining of a gel should also be conducted carefully to prevent protein diffusion. Later is one example (all procedures can be carried out at room temperature): Step 1. Fixation: gently shake a gel in a solution containing 40% MeOH and 10% acetate for 30 min. Step 2. Staining: gently shake the gel in a solution containing 0.025% CBB G-250 and 10% acetate for 60 min. Step 3. Destaining: wash the gel in 10% acetate with gentle shaking for 15 min 3  and more (sometimes it is better to destain a gel in water overnight).

3.2 Cleavage of Synthetic SPs With Small Tags In Vitro An example of an in vitro cleavage assay using synthetic peptides is shown in Fig. 4. The synthetic Myc-SPBla-Flag peptide was cleaved by purified RsePHis6-Myc to generate cleavage products (F1 and F2 in Fig. 4B). A typical protocol for an in vitro cleavage reaction is as follows: this is a slight modification of the reaction described in Section 2.2.2. 1. Mix a chemically synthesized target peptide (150 μg/mL) with the purified enzyme (30 μg/mL) in 1  RseP buffer (see Section 2.2.2) supplemented with 100 mM NaCl and 1  protease inhibitor cocktail (EDTA-free) (Nacalai Tesque, Inc., catalog #: 03969). 2. Incubate samples at 37°C. Withdraw aliquots at certain intervals and mix them with an equal volume of appropriate 2  SDS sample buffer (for the PAGE system used) to stop the reaction.

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Y. Hizukuri et al.

3. Incubate the samples at 85°C for 2 min (for analysis by means of the tricine SDS-PAGE system) or at 70°C for 5 min (for the Bis-Tris SDS-PAGE system) for denaturation of proteins. 4. Proteins are separated by an appropriate gel system and detected by CBB G-250 staining with the precautions described in Section 3.1.

3.3 Proteolysis of Authentic SPs Inside the Membrane The experimental system for in vitro translocation of a secretory protein across the bacterial membranes has been established (Mizushima, Tokuda, & Matsuyama, 1991) (Fig. 5A). In this system, an in vitrosynthesized precursor form of a secretory protein (containing an SP at the N-terminus) is translocated into the IMVs prepared from the bacterial cytoplasmic membrane, resulting in accumulation of the remnant SP in the membrane. We have utilized this translocation system to prepare a membrane-integrated authentic SP and examined its degradation by RseP in the membrane. An example of the results is shown in Fig. 5B and C. IMVs were prepared from a WT strain carrying no plasmids or from a strain overexpressing the WT RseP (+RseP) or the protease-inactive mutant (+H22F). The IMVs were mixed with a precursor form of an in vitro-synthesized [35S]Metlabeled model secretory protein, LivK-Lpp. LivK-Lpp is a chimeric protein in which the LivK SP region is followed by a small Lpp (lipoprotein) mature sequence (Fig. 5B). We used this model protein because we found that LivK SP can be efficiently cleaved by RseP in our in vivo assay system (Saito et al., 2011), and previous studies showed that a model protein with a similar construct (such as OmpF-Lpp) showed high efficiency of translocation into IMVs in vitro (Hikita & Mizushima, 1992). Immediately after the mixing, LivK-Lpp was translocated across the membrane, and the remnant SP was generated. The SP signal decreased gradually during further incubation when IMVs from the WT strain were used, indicating that the SP was degraded. Overproduction of RseP, but not that of the H22F mutant, enhanced the degradation (Fig. 5C). These results showed that RseP can degrade an authentic SP. Typical protocols for the IMV preparation and the in vitro synthesis of a model secretory protein are described in the following sections. 3.3.1 Preparation of IMVs for In Vitro Translocation 1. Grow cells expressing a target protein at 30°C in 1 L of the L medium until the mid-log phase.

A 3. In vitro translocation & SP proteolysis (Pe

ripl

asm

)

(Cy

1 Translocation

Secreted protein

Lep

2

3

top

las

m)

Sec Translocon

RseP

SP IMV

E. coli cell

(Cytoplasm) IM

Periplasm Cytoplasm

IM

PG

LivK-Lpp

SP

(Periplasm)

Mix OM

2. In vitro transcription & translation

1. Preparation of IMVs B

LivK-Lpp (SPLivK-Lpp) Lep MKRNAKTIIAGMIALAISHTAMA DESSN--

LivK 1

C

2

3

4

Lpp 5

6

7

8

9

Precursor mature

10 5 2

SP Time (min)

IMV

(kDa)

0

4

WT

12

0

4

+RseP

12

0

4

12

+H22F

Fig. 5 Proteolysis of an authentic SP by RseP in the inverted membrane vesicles (IMVs). (A) Schematic representation of the assay. Step 1: IMVs are prepared from wild-type (WT) or RseP-overexpressing cells. Step 2: The [35S]Met-labeled model presecretory protein (LivK-Lpp) is synthetized by in vitro transcription/translation. Step 3: The synthesized presecretory proteins are mixed with IMVs to allow for translocation across the membrane. SPs generated by Lep-catalyzed processing are degraded by RseP. (B) The amino acid sequence of the N-terminal region of the model presecretory protein LivK-Lpp, which contains the SP region of LivK followed the Lpp mature sequence. (C) Detection of the authentic SP of LivK-Lpp generated upon in vitro translocation of the model protein into IMVs. Proteins were separated by means of the tricine SDS-PAGE system (10–20% gradient gel). IMVs prepared from the WT cells (lanes 1–3), from cells overproducing the WT RseP (lanes 4–6), or from cells overproducing the active-site mutant (lanes 7–9) of RseP were used. Panels (B) and (C) are modified from Saito, A., Hizukuri, Y., Matsuo, E., Chiba, S., Mori, H., Nishimura, O., et al. (2011). Postliberation cleavage of signal peptides is catalyzed by the site-2 protease (S2P) in bacteria. Proceedings of the National Academy of Sciences of the United States of America, 108(33), 13740–13745.

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2. Harvest cells by centrifugation, wash them with 20 mL of 10 mM Tris– HCl (pH 8.1), centrifuge again, and resuspend in a mixture of 30 mL of spheroplast buffer (30 mM Tris–HCl (pH 8.1) and 20% sucrose) and 3 mL of 1 mg/mL lysozyme in 0.1 M EDTA (pH 8.0). 3. Incubate the cells at 0°C for 30 min to convert them to spheroplasts, collect by centrifugation, and resuspend in 15 mL of a buffer consisting of 10% sucrose, 3 mM EDTA (pH 7.0), and 1 mM Pefabloc SC (MilliporeSigma (MerckMillipore), Darmstadt, Germany, catalog #: 124839). 4. Disrupt spheroplasts by French press (6000 psi  2 ), remove cell debris by low-speed centrifugation (3600  g, 10 min at 4°C), and recover the total membrane fraction by ultracentrifugation (150,000  g, 30 min at 4°C). Cell disruption using French press converts the inner membrane (IM) to IMVs. 5. Resuspend the collected membranes in 1.5 mL of 3 mM EDTA (pH 7.0) with 1 mM DTT by repeated passages through a 26-gauge needle with a 2.5-mL syringe. 6. Place 0.8 mL of the sample on top of two layers of sucrose-EDTA solutions (1.5 mL of 33% sucrose with 3 mM EDTA; 2.5 mL of 48% sucrose with 3 mM EDTA) in a 5-mL centrifuge tube, centrifuge the samples at 120,000  g for 16 h at 4°C (with a swing rotor), and then collect the IM fraction from the boundary between the 33% and 48% sucrose layers. 7. Dilute the membranes with five volumes of buffer E0 -glycerol (50 mM Hepes–KOH (pH 7.5), 50 mM KCl, and 5 mM magnesium acetate), precipitate them by ultracentrifugation (180,000  g, 30 min at 4°C), and resuspend them in 200 μL of a buffer consisting of 50 mM Hepes– KOH (pH 7.5), 10% glycerol, and 8.5% sucrose as described earlier. 8. Divide the prepared IMV samples into aliquots, freeze them in liquid nitrogen, and store at –80°C. 3.3.2 Preparation of a Model Secretory Protein for In Vitro Translocation/SP Cleavage Assay A protocol for mRNA preparation: 1. Insert a gene for the model secretory protein under control of the SP6 promoter in a vector (such as pSP65; Melton et al., 1984). 2. Linearize the plasmid DNA by digestion with an appropriate restriction enzyme, purify it by phenol/chloroform extraction followed by ethanol precipitation, and dissolve in H2O.

Functional and Structural Analysis of RseP

17

3. Incubate the linearized template DNA (5 μg) in a buffer (consisting of 40 mM Tris–HCl (pH 7.5), 6 mM MgCl2, 2 mM spermidine, 10 mM DTT, and 0.01% BSA; supplied as 10 buffer by Takara Bio, Inc., Shiga, Japan) with 0.5 mM each NTP, RNase inhibitor (trace amount), and SP6 RNA polymerase (30 U/μL; Takara Bio, Inc., catalog #: 2520A) at 40°C (or 37°C) for 2 h. The reaction mixture can be used as “mRNA” for the following in vitro protein synthesis and stored at –80°C. A protocol for preparation of a [35S]Met-labeled model presecretory protein: 1. Mix appropriate amounts of model substrate-encoding mRNAs with the S100 fraction (containing ribosomes), [35S]Met, and small molecules including amino acids, nucleotides, salts, and other reagents and incubate the mixture at 30°C for 40 min. See Mizushima et al. (1991) for details of the in vitro protein synthesis with the S100 fraction. 2. Translated proteins are precipitated by mixing the reaction mixture with a 10-fold volume of 5% TCA, collected by centrifugation, washed with 5% TCA, washed again with acetone, and finally dissolved in a buffer consisting of 50 mM Hepes–KOH (pH 7.5) and 6 M urea. You can store urea-denatured proteins at –20°C. Urea denaturation is required for efficient translocation of the model protein into IMVs. 3.3.3 Translocation of an In Vitro-Synthesized and [35S]Met-Labeled Substrate Protein Into IMVs and Analysis of SP Proteolysis by RseP Protocols for translocation and analysis of substrate cleavage (here, LivK-SP) by RseP are described later: 1. Incubate IMVs in a buffer consisting of 50 mM Hepes–KOH (pH 7.5), 5 mM MgSO4, 19.6 μg/mL purified SecA, 19.2 μg/mL purified SecB, 5 mM sodium succinate, 5 mM ATP, and an appropriate amount of a urea-denatured substrate (e.g., LivK-Lpp) at 30°C (for purification of SecA and SecB, see Mizushima et al., 1991). We usually make the final urea concentration in the reaction mixture lower than 0.3 M because a high concentration of urea may interfere with the subsequent reactions. Because the protein translocation and the accompanying SP formation are rapid events, the reactions are best initiated by addition of the substrates to a prewarmed reaction mixture without substrates.

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Y. Hizukuri et al.

2. After incubation at 30°C for 3 min, add NaN3 (final conc. 50 mM), NaCl (final conc. 1 M), and zinc acetate (final conc. 50 μM) to stop translocation and to stimulate SP degradation. 3. Continue incubation at 30°C to chase SP degradation. 4. Withdraw samples at appropriate time points, mix them with a 10-fold volume of 5% TCA, and incubate at 0°C to precipitate proteins. Collect TCA-precipitated samples, wash with acetone, and dissolve in the appropriate 1 sample buffer (e.g., tricine or LDS sample buffer) by vigorous mixing at room temperature for 30 min followed by heating at 37°C for 5 min. 5. Analyze SPs and their cleavage products by means of an appropriate SDS-PAGE system for low-molecular-weight proteins (see Section 3.1).

4. STRUCTURAL ANALYSIS OF RseP USING THIOL-SPECIFIC MODIFICATION REAGENTS Modifiability of engineered Cys residues by membrane-permeable or impermeable thiol-specific alkylating reagents provides information on local structures of a membrane protein especially from the standpoint of interaction with a membrane. Here, we show several examples of application of this approach to analysis of RseP. The results described in Sections 4.1 and 4.2 reveal that the thiol modification assays using various reagents are useful for evaluation of the structure of (and an environment around) membrane protein domains. Structural information on a membrane protein in its native state would be important for elucidation of its physiological functions.

4.1 Evaluation of the Environment Around the Protease Active-Site Region of RseP I-CLiPs including RseP are believed to catalyze proteolysis of membrane proteins within the lipid bilayer and indeed have their active-site residues in predicted transmembrane regions. Thus, the active site of the I-CLiPs should be accessible to a water molecule that is required for hydrolysis of a peptide bond, while it is embedded in the hydrophobic membrane. To biochemically explore the environment around the active-site regions of RseP, we examined the accessibility of a membrane-impermeable thiol-alkylating reagent (4-acetamido-40 -maleimidylstilbene-2,20 -disulfonic acid, AMS) (Fig. 6A) to Cys residues introduced into the active-site region

19

Functional and Structural Analysis of RseP

A IMV mal-PEG-5k

1st: AMSlabeled

IM

(M.W.= ~5,000)

2nd: malPEG-unlabeled

SH

RseP

AMS (M.W.= 536.44)

N

(PDZ-C)

1st: AMSunlabeled

(PDZ-N)

C

NEM

2nd: malPEG-labeled (band upshift)

(M.W.= 125.13)

SH

V13

A77

E23

B

(%) 100 80 60 40 20 0

(%) 100 80 60 40 20 0

V13C

0

60 50 -

Time(min) 0 5 15 0 5 15 0 5 15 Triton X-100 – + – Gdn + + –

αMyc

: + Gdn

AMS modification

60 -

A77C 50 -

AMS modification

(kDa) 1 2 3 4 5 6 7 8 9 60 V13C 50 -

E23C

: no addition

RseP-His6-Myc

AMS modification

mal PEG modified

5

15 (min)

: + Gdn/ TX

(%) 100 80 60 40 20 0

A77C

0

5

15 (min)

E23C

0

5

15 (min)

Fig. 6 AMS modifiability of the Cys residues introduced into RseP. (A) Schematic diagrams explaining the two-step AMS/mal-PEG-5k modification assay. Information about some thiol-alkylating regents is shown in the left. (B, left half ) Immunological detection of the mal-PEG-5k-modified RseP-His6-Myc. The membrane samples prepared from the cells expressing the indicated single-Cys derivatives of RseP-His6-Myc were used. (Right half ) The proportion of the AMS-modified single-Cys derivatives of RseP-His6-Myc was calculated as the ratio of the mal-PEG-5k-unmodified to the total RseP-His6-Myc and then depicted graphically. Two independent experiments were carried out, and the mean values are shown with the standard deviations.

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Y. Hizukuri et al.

of RseP (Koide, Maegawa, Ito, & Akiyama, 2007; Maegawa, Koide, Ito, & Akiyama, 2007). If the introduced Cys residue is sufficiently exposed to the external milieu, then it should be easily modified with AMS. In contrast, if the Cys residue faces the hydrophobic core region of the membrane or is located inside the folded protein domain, then it should be unmodifiable by AMS. On the other hand, membrane solubilization with a detergent (Triton X-100) or protein unfolding with a denaturant (guanidine hydrochloride; Gdn-HCl), respectively, should make the Cys residue modifiable with AMS in each case. Because AMS is a small molecule (536 Da), it is often difficult to detect AMS modification of a target protein by changes in mobility during standard SDS-PAGE. The second modification with methoxypolyethylene glycol 5000 maleimide (mal-PEG-5k), a thiol alkylating regent of 5 kDa, enables definitive discrimination of the AMS modification; when AMS-treated samples are next treated with mal-PEG-5k after SDS denaturation of proteins, only free Cys residues that did not undergo prior modification by AMS will react with mal-PEG-5k, and mal-PEG modification can be easily detected as a significant upshift of the modified bands during SDS-PAGE. This method is suitable for evaluation of the fraction of the AMS-modified species in the total target protein and thus is useful for kinetic analyses of the modification. Typical results of this assay are shown in Fig. 6B. The Val13 residue of RseP is located in the middle of the first transmembrane segment, and Ala77 is in the first cytoplasmic loop region of RseP. The V13C substitution was modified by AMS only in the presence of both Gdn-HCl and Triton X-100. In contrast, the A77C substitution was efficiently modified with AMS even in the absence of Gdn-HCl and Triton X-100. These results strongly indicate that Val13 is embedded in the lipid phase of the membrane, whereas Ala77 is located on the surface of a soluble domain. Glu23 is a catalytic residue of RseP, which should coordinate a water molecule for the hydrolytic reaction. Cys introduced at Glu23 was not modified by AMS when the membranes were directly treated with AMS in the absence of Gdn-HCl and Triton X-100 but was fully modified in the presence of both reagents. In the presence of Gdn-HCl only, E23C was partially modified by AMS. These results suggest that Glu23 is not completely embedded in the lipid phase but is located within a folded protein domain that is partially exposed to an aqueous environment. The following is a typical protocol for the AMS modification assay: 1. Grow cells expressing a single-cysteine derivative of a target protein at 30°C in 5 mL of the L medium until the early- to mid-log phase.

Functional and Structural Analysis of RseP

21

2. Harvest cells by centrifugation, wash with 10 mM Tris–HCl (pH 8.1), resuspend them in a buffer consisting of 10 mM Tris–HCl (pH 8.1) and 1 mM PMSF, and sonically disrupt at 0°C. 3. After removal of unbroken cells by low-speed centrifugation, ultracentrifuge the cleared lysates (100,000  g, 60 min at 4°C) and dissolve the membrane pellets in a buffer consisting of 50 mM Tris–HCl (pH 8.1), 15% glycerol, and 1 mM Tris(2-carboxyethyl)phosphine (TCEP; SigmaAldrich Co. (Fluka), MilliporeSigma (Sigma-Aldrich (Fluka)), Darmstadt, Germany, catalog #: C4706). TCEP, a reducing agent, is added to keep the side chain of the introduced single-Cys residue in a reduced form. TCEP is useful in experiments using maleimide derivatives because it does not interfere with modification of cysteine residues by maleimide, unlike other generally used reductants such as DTT and ME. 4. Divide the membrane samples into three portions (9 μL each), mix with 1 μL of water (samples A and B) or 10% Triton X-100 (sample C), and incubate for 30 min on ice followed by 5 min at 24°C. 5. Add 20 μL of a buffer consisting of 37.5 mM Tris–HCl (pH 8.1) and 1.5 mM AMS (Thermo Fisher Scientific Inc. (Invitrogen (Molecular Probes)), Waltham, MA, catalog #: A485) to sample A; 20 μL of a buffer consisting of 37.5 mM Tris–HCl (pH 8.1), 1.5 mM AMS, and 6 M Gdn-HCl to sample B; and 20 μL of a buffer consisting of 37.5 mM Tris–HCl (pH 8.1), 1.5 mM AMS, 6 M GdnHCl, and 1.5% Triton X-100 to sample C and incubate them at 24°C for appropriate periods. 6. Terminate the modification reaction by addition of 2 μL of 1 M DTT (final conc. 62.5 mM) followed by incubation at 24°C for 18 min. 7. The total proteins are precipitated with TCA, washed with 5% TCA, and then washed again with acetone. 8. Solubilize the proteins with 18 μL of a buffer consisting of 100 mM Tris– HCl (pH 8.1), 1% SDS, and 1 mM TCEP with vigorous mixing at room temperature for 30 min followed by incubation at 37°C for 5 min. 9. After addition of 2 μL of 50 mM mal-PEG-5k (MilliporeSigma (Sigma-Aldrich (Fluka)), Darmstadt, Germany, catalog #: 63187; dissolved in dimethyl sulfoxide (DMSO)), the samples are vigorously mixed at 37°C for 30 min. Note that in reactions of modification by mal-PEG, the optimal temperature (4–37°C), reaction duration (a few minutes to 1 h), and mal-PEG concentration (1–5 mM) may depend on an individual experiment, and thus you should customize the conditions.

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10. The samples are mixed with 2  SDS sample buffer, incubated at 37°C for 5 min, and analyzed by SDS-PAGE in a 7.5% gel with immunoblotting.

4.2 Evaluation of Solvent Accessibility of the Periplasmic Domain of RseP The Cys modifiability assay can also be employed to probe structural features of the periplasmic domains of a target membrane protein using spheroplasts in which increased permeability of the outer membrane allows for entry of chemical modifiers into the periplasmic space. This assay will be especially useful when the three-dimensional structural information of a target protein (or a domain) is available. Modifiability of Cys residues (introduced at various positions) with thiol-alkylating reagents will shed light on protein– protein and protein–membrane interactions because these interactions will interfere with modification of an engineered Cys residue as described in the previous section. Furthermore, comparison of the accessibility of an engineered Cys residue to thiol-alkylating reagents of different sizes (e.g., mal-PEG-5k, 5 kDa; AMS, 536 Da; n-ethyl-maleimide (NEM), 125 Da) will give clues to local structures and disposition of the region around the Cys-modified site. For example, if the target site is accessible to AMS but not to mal-PEG-5k, then this site is likely to be exposed to the external milieu but located at a recessed position such that access of a large molecule is limited by protein structures and/or the membrane. Accessibility of thiol-alkylating reagents with a different membrane permeability (for example, NEM is membrane-permeable, but mal-PEG-5k and AMS are not) will provide insights into the interaction of the Cys-modified site with the membrane. RseP has the periplasmic region containing two tandemly arranged PDZ domains (PDZ tandem). An X-ray crystallographic study of the isolated PDZ tandem showed that it has a clam-like structure with a narrow cavity named the PDZ pocket (Hizukuri et al., 2014). To analyze the structure of the PDZ tandem in intact, membrane-embedded RseP, we have performed the mal-PEG accessibility assay based on the information from the X-ray crystallographic structure of the PDZ tandem. An example of the results of mal-PEG-5k modification assays is shown in Fig. 7. The Cys residue introduced into the position expected to be located inside the PDZ pocket (V261C) showed no detectable mal-PEG-5k modification, whereas the one introduced at the position expected to be located on the external surface of the PDZ domain (A136C) was modified rapidly. V261C was modified with

23

Functional and Structural Analysis of RseP

Outside PDZ pocket

Inside PDZ pocket

A

PDZ-C

A136

V261

PDZ-N

PDZ pocket

C

D163

TM2

3-C

Zn2+

Proximal to membrane

TM1

TM4

3-N

N

RseP B (RseP) (Triton X-100)

A136C –

+

V261C –

+

Time (min) 0 1 5 0 1 5 0 1 5 0 1 5 80 60 50 -

malPEG -labeled * unlabeled

40 (RseP) (Triton X-100)

D163C –

+

Cys-less –

+

Time (min) 0 1 5 0 1 5 0 1 5 0 1 5 80 -

malPEG -labeled * unlabeled

60 50 40 1 2 3 4 5 6 1 2 3 4 5 6

αMyc

Fig. 7 The mal-PEG-5k modification assay for the periplasmic domain of the RseP. (A) Structural model of full-length RseP and the residues replaced with Cys around the PDZ pocket structure. (B) Results of the modification assay with spheroplasts. The bands of the mal-PEG-5k-modified RseP proteins are significantly upshifted in the SDSpolyacrylamide gel. Modified from Hizukuri, Y., Oda, T., Tabata, S., Tamura-Kawakami, K., Oi, R., Sato, M., et al. (2014). A structure-based model of substrate discrimination by a noncanonical PDZ tandem in the intramembrane-cleaving protease RseP. Structure, 22(2), 326–336.

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Y. Hizukuri et al.

mal-PEG-5k under detergent solubilization conditions, possibly because the membrane solubilization alters the conformation of the PDZ tandem slightly, and this situation may enable the access of mal-PEG-5k to the interior of the PDZ pocket. Because V261C was modifiable with the smaller reagent AMS even in the absence of a detergent, the interior of the PDZ pocket is open to the external milieu, but the entry site is not large enough for a bulky reagent such as mal-PEG-5k to pass through when RseP is embedded in the membrane (Hizukuri et al., 2014). D163C was not modified with mal-PEG-5k in the absence of a detergent, even though this Cys residue is expected to be located on the external surface of the PDZ domain. Nonetheless, D163C became modifiable when the membrane was solubilized, suggesting that this residue is located proximally to the membrane surface and is inaccessible to the reagent due to steric hindrance from the membrane. A typical protocol (a slightly modified version of the one in Hizukuri et al., 2014) for the mal-PEG-5k modification assay using spheroplasts is as follows. Modification with other thiol-alkylating reagents can be carried out essentially in the same way: 1. Grow cells expressing a single-cysteine derivative of a target protein at 30°C in 5 mL of the L medium until the early- to mid-log phase. 2. Convert the cells to spheroplasts by lysozyme/EDTA treatment in spheroplast buffer (as described in Section 3.3.1). The sucrose concentration (20%) should be kept constant throughout the reaction steps because sucrose dilution sometimes leads to bursting of spheroplasts, thus making the results unreliable. 3. Mix the spheroplasts with 1/200 volume of MgCl2 (final conc. 5 mM), 1/100 volume of PMSF (final conc. 1 mM), and 1/500 volume of TCEP (final conc. 1 mM). 4. Incubate the spheroplast samples with (or without) 2% Triton X-100 at 0°C for 3 min to lyse (or not lyse) them. 5. Mix each sample with an equal volume of 2  reaction buffer (60 mM Tris–HCl (pH 8.1), 20% sucrose, and 2 mM mal-PEG-5k) and incubate the mixture for a few minutes at 4°C. We usually use 50 mM mal-PEG5k in DMSO as a stock solution to prepare 2  reaction buffer. 6. Stop the reaction by adding 10% ME. Proteins are then precipitated with TCA and analyzed by SDS-PAGE and immunoblotting. Because a high concentration of sucrose may interfere with TCA precipitation, it is recommended to dilute sucrose before TCA precipitation or to add small

Functional and Structural Analysis of RseP

25

volumes of 20% Triton X-100 to the reaction mixture as a carrier to facilitate the precipitation.

4.3 Determination of In Vivo Substrate Cleavage Sites Using mal-PEG Modification We will describe another application that involves mal-PEG modification. We used a combination of cysteine-scanning mutagenesis and mal-PEG modification to determine cleavage sites of the RseP substrates within the membrane. In this assay, a Cys residue is introduced into various positions in the transmembrane region of substrates. After RseP-catalyzed in vivo cleavage of these substrates, N-terminal cleavage products are examined to test whether they contain the introduced Cys residue. The presence of the Cys residue can be assessed by modifiability with mal-PEG (Fig. 8A). Our results indicated that RseP cleaves HA-MBP-RseA140 between Ala108 and Cys109 within the membrane (Akiyama et al., 2004). Consistent results were obtained when we carried out the mal-PEG-modification assay using purified preparations of RseP and a model substrate (His6MBP-RseA140) (Akiyama et al., 2004). Furthermore, analysis of in vivoaccumulated RseA cleavage products confirmed the cleavage site (Flynn, Levchenko, Sauer, & Baker, 2004), thus supporting effectiveness of this approach. It should be noted that a Cys substitution may significantly interfere with cleavage and/or shift the site of cleavage in some cases. Therefore, this approach seems to be useful to at least roughly estimate the site of an intramembrane cleavage in a living cell. The typical results are shown in Fig. 8. Because the transmembrane region of RseA originally contains a Cys residue (Cys109), which makes interpretation of the results complicated, here we show the data with another model substrate (HA-MBP-RseA(LY1)140). The modification by mal-PEG caused an almost complete upshift for the N-terminal cleavage product (CL) of HA-MBP-RseA(LY1/F12C)140 but not for HA-MBPRseA(LY1/F17C)140, indicating that the cleavage occurred between Phe12 and Phe17. Detailed analyses revealed that RseP mainly cleaved HA-MBP-RseA(LY1)140 between Phe16 and Phe17 with a minor cleavage between Phe15 and Phe16. The following is a typical protocol: 1. Grow cells expressing a single-cysteine derivative of a target protein at 30°C in the L medium until the early- to mid-log phase.

26

Y. Hizukuri et al.

A HA-MBPRseA(LY1)140

C C

CL

UC

RseP

MBP

MBP

HA

HA

N

rseP +

F12C

ΔrseP

C

N

rseP +

F17C

ΔrseP

C malPEG

C

C malPEG

17 SH 17 SH

malPEG

malPEG 12 SH

12 SH

N

N

(CL)-malPEG

N

(UC)-malPEG

N (CL)

(UC)-malPEG

B

UC-malPEG CL-malPEG UC CL malPEG – + – + – + – + rseP + + Δ Δ WT(Cysless)

F12C

– + – + + Δ F17C

αHA

Fig. 8 The in vivo approach to the determination of RseP cleavage sites in a model substrate. (A) Schematic explanation of the assay with derivatives of model substrate HA-MBP-RseA(LY1)140 shown in (B). (B) Application to the determination of the cleavage site in HA-MBP-RseA(LY1)140. The open vertical arrows indicate the sites of the RseP cleavage. Modified from Akiyama, Y., Kanehara, K., & Ito, K. (2004). RseP (YaeL), an Escherichia coli RIP protease, cleaves transmembrane sequences. EMBO Journal, 23(22), 4434–4442.

2. Whole-cell proteins are then precipitated by TCA treatment and solubilized in a buffer consisting of 100 mM Tris–HCl (pH 8.1), 1% SDS, and 1 mM TCEP by vigorous mixing at room temperature for 30 min. 3. Add 1/10 volume of 50 mM mal-PEG-5k (in DMSO) to the samples and incubate them further at room temperature for 30 min with

Functional and Structural Analysis of RseP

27

vigorous mixing. (The mal-PEG treatment conditions should be tuned in each experiment (see Section 4.1). In case the efficiency of modification by mal-PEG is low, protein solubilization and mal-PEG modification can be conducted at 37°C. 4. Mix the samples with an equal volume of 2  SDS sample buffer (containing ME; final conc. 10%), incubate them at 37°C for 5 min, and analyze by SDS-PAGE and immunoblotting (free mal-PEG is quenched by ME in SDS sample buffer).

5. IN VIVO CROSS-LINKING ANALYSIS TO PROBE THE INTERACTION OF RseP WITH ITS SUBSTRATE IN THE MEMBRANE I-CLiPs interact with their substrates in the membrane. A crosslinking approach is a powerful tool for analysis of in vivo interactions between proteins and can be applied to intramembrane protein interactions. In this part of the chapter, we show two applications of in vivo cross-linking approaches to analysis of the RseP–substrate interaction.

5.1 In Vivo Photo Cross-Linking This method is a rather new technique to probe protein interactions in living cells (Akiyama et al., 2015; Davis & Chin, 2012). In this method, a photoreactive nonnatural amino acid such as p-benzoyl-L-phenylalanine (pBPA) is introduced into essentially any desired positions in a target protein using an amber suppression mechanism (Fig. 9A). This method provides a means to probe not only stable but also transient protein–protein interactions. After ultraviolet (UV) light irradiation of cells expressing a pBPA-containing protein, pBPA forms a covalent bond with a CH group of a neighboring molecule. Cross-linked partner proteins can be identified immunologically or by mass spectrometry, although determination of the exact site of cross-linking in the cross-linked partner protein is often difficult. Using this approach, we analyzed the interaction between plasmidexpressed RseP-His6-Myc and chromosomally encoded RseA (Akiyama et al., 2015) (Fig. 9B and C). We used a strain with deletions of ompA, ompC, and rseP as a host because rseP can be deleted even in an rseA+ background in the absence of the ompA and ompC genes. Plasmid pEVOL-pBpF (Young, Ahmad, Yin, & Schultz, 2010), which encodes an evolved amber suppressor tRNA and tyrosyl-tRNA synthetase pair, allows for incorporation of pBPA into an amber codon site. Cells harboring pEVOL-pBpF were further

28

Y. Hizukuri et al.

A

Step. 1:

Step. 2:

incorporation of pBPA

photo cross-linking

Cell culture

Step. 1

Step. 2

Plasmid (pEVOL-pBpF)

UV irradiation (365 nm)

Amber suppressor tRNA

Evolved tyrosyl-tRNA synthetase

pBPA (photocrosslinker) Binding partners neighboring molecules

mRNA 3′ …… UAG ……

5′

pBPAincorporated protein

TAG …

Covalently cross-linked

Plasmid (target protein with amber mutation)

B

C RseP

(PDZ)

N

C

αRseA

C

(kDa) 82 68 55 42

1

2

3

4

5 6 XL

RseA

28

RseA148

15

αMyc

120 100

V70

Y69

σE N

RseA

80 60 50

40 UV irrad. pBPA

RseP-His6-Myc

+ – + – + – Y69

V70

none

Fig. 9 In vivo photo cross-linking between RseP and RseA. (A) An outline of the amber suppression-mediated site-specific introduction of pBPA into the target protein (Step 1) and formation of a cross-linked product after UV irradiation (Step 2). (B) Schematic explanation of the results shown in (C). (C) Immunological detection of the cross-linked product by an anti-RseA (upper panel) or anti-Myc antibody (lower panel). The RseP derivative with an amber mutation at the indicated position was expressed in the cells harboring pEVOL-pBpF. RseA, full-length RseA; RseA148, DegS-cleaved RseA; XL, crosslinked product. Modified from Akiyama, K., Mizuno, S., Hizukuri, Y., Mori, H., Nogi, T., & Akiyama, Y. (2015). Roles of the membrane-reentrant beta-hairpin-like loop of RseP protease in selective substrate cleavage. Elife, 4, e08928.

Functional and Structural Analysis of RseP

29

transformed with a plasmid (a pUC-based vector; expression of the cloned gene was under control of the lac promoter) encoding a variant of RsePHis6-Myc carrying an amber mutation at position Tyr69 or Val70. Both RseP mutants additionally carried a mutation of a catalytic residue (E23Q) to prevent substrate cleavage. The RseP(Y69pBPA)-His6-Myc mutant generated a band corresponding to the RseP-RseA cross-linked product that was detected with an anti-RseA antibody in a UV irradiation-dependent manner (Fig. 9C, lanes 1 and 2). In contrast, no such band was observed with RseP(V70pBPA)His6-Myc or pBPA-free RseP-His6-Myc. These results indicate that residue Tyr69, but not Val70, of RseP is in close proximity to RseA (Fig. 9B). 1. Grow cells carrying pEVOL-pBpF and a plasmid for a target protein at 30°C in the M9 medium supplemented with 0.02% arabinose (for induction of the evolved tyrosyl-tRNA synthetase) and 1 mM pBPA (Bachem AG, Bubendorf, Switzerland, catalog #: 4017646) until the early- or mid-log phase. 2. Cell cultures are chilled on ice, and spectinomycin (final conc. 100 μg/ mL) is added to stop further protein synthesis (note that pEVOL-pBpF carries a chloramphenicol resistance marker gene). 3. Remove a portion of the culture and irradiate it with UV light (365 nm) at 4°C for 10 min in a Petri dish by means of a B-100AP UV lamp (UVP, LCC, Upland, CA) at a distance of 4 cm. UV-irradiated cells are then mixed with 1/20 volume of 100% TCA. 4. The precipitated proteins are collected by centrifugation, washed with acetone, and dissolved in 1  SDS sample buffer with vigorous mixing at room temperature for 30 min, followed by incubation at 37°C for 5 min and analysis by SDS-PAGE in a 10% gel with immunoblotting.

5.2 Disulfide Cross-Linking In contrast to the in vivo cross-linking method described earlier, the disulfide cross-linking approach is generally used to analyze a possible interaction between two defined sites. A Cys replacement is introduced into each of these sites, and disulfide bond formation between the introduced Cys residues is examined. We applied this approach to analysis of the RseP–substrate interaction (Akiyama et al., 2015; Koide, Ito, & Akiyama, 2008). In this experiment, single-Cys derivatives of RseP-His6-Myc (additionally carrying the E23Q active-site mutation) and HA-RseA140 were coexpressed from two compatible plasmids (a pUC-based vector was used for RseP, and a

30

Y. Hizukuri et al.

pACYC184-based vector for RseA) in a strain with deletions of rseA, rseP, and degS. When RseP(Y69C)-His6-Myc and HA-RseA(A108C or C109) 140 were coexpressed, a cross-linked product was generated in an oxidantdependent manner (Fig. 10A, lanes 4 and 6). These bands were not detected in the samples treated with ME. Coexpression of either RseP(V70C)-His6Myc or RseP(Cys-less)-His6-Myc yielded no cross-linked product. These results suggest that residue Tyr69, but not Val70, of RseP is in close proximity to RseA-TM; this finding matches the results of the in vivo photo cross-linking experiments (see Section 5.1). 1. Grow cells expressing a pair of single-Cys derivatives of RseP and a model substrate at 30°C in the L medium until the early- or mid-log phase. 2. Cell cultures are chilled on ice, and chloramphenicol (final conc. 200 μg/mL) is added to stop further protein synthesis. 3. Wash the cells with 10 mM Tris–HCl (pH 8.1) and resuspend them in the same buffer. 4. Disulfide bond formation is induced by addition of the oxidative reagent Cu2+(phenanthroline)3 (final conc. 0.1 mM; this reagent should be freshly prepared by mixing one volume of 240 mM CuSO4 with two volumes of 360 mM PT) with subsequent incubation at 37°C for 5 min. Another portion is treated with 3 mM PT as a no-oxidation control. 5. Terminate the oxidation reaction by adding neocuproine (Nacalai Tesque, Inc., catalog #: 24115; final conc. 12.5 mM; it competes with PT for chelating a copper ion and quenches the oxidizing reaction) followed by incubation at 37°C for 5 min. 6. Proteins are precipitated with 5% TCA, washed with acetone, and dissolved in a buffer consisting of 100 mM Tris–HCl (pH 7.5), 1.5% SDS, and 5 mM EDTA. 7. The samples are divided into two portions. One portion (nonreduced condition) is mixed with NEM (final conc. 25 mM) to block free Cys residues and then with an equal volume of 2 SDS sample buffer without the reducing reagent. The other portion (reduced condition) is mixed with water instead of NEM and then with 2 SDS sample buffer (containing 20% ME, final conc. 10%). Both samples are heated at 98°C for 5 min and analyzed by SDS-PAGE in a 10% gel with immunoblotting.

31

Functional and Structural Analysis of RseP

A

No ME

α HA

(kDa) 150 −

HA-RseA140

RseP-His6-Myc

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24

75 − 50 − 37 −

XL

20 −

α Myc

150 − 75 − 50 − 37 −

XL

Y69C

r

−+ − + − +− + Vect o

Vect o

Vect o

RseA RseP

r

r

Cu2+(PT)3 − + − + − + − + − + − + − + − +

V70C

Cys-less

B

α HA

+ ME (kDa) 150 − 75 − 50 − 37 −

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24

20 − 75 − 50 − 37 −

RseP

Y69C

r

Vect o

r Vect o

RseA

− + −+ −+ −+

r

Cu2+(PT)3 − + − + − + − + − + − + − + − +

Vect o

α Myc

150 −

V70C

Cys-less

Fig. 10 Disulfide cross-linking between RseP and RseA. Immunological detection of the cross-linked product by an anti-HA (upper panels) or anti-Myc antibody (lower panels). The samples were treated without (A) or with (B) 2-mercaptoethanol (ME). Cys-less or the indicated single-Cys derivatives of RseP-His6-Myc and HA-RseA140 were coexpressed in the cells. Asterisk, a possible dimer of RseP-His6-Myc; XL, cross-linked product. Modified from Akiyama, K., Mizuno, S., Hizukuri, Y., Mori, H., Nogi, T., & Akiyama, Y. (2015). Roles of the membrane-reentrant beta-hairpin-like loop of RseP protease in selective substrate cleavage. Elife, 4, e08928.

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ACKNOWLEDGMENTS We thank Dr. Hiroyuki Mori and other members in our laboratory for helpful discussions. This work was supported by JSPS KAKENHI (Grant Numbers 26840033 (to Y.H.) and 15H04350 (to Y.A.)) and MEXT KAKENHI (Grant Number 15H01532 (to Y.A.)).

REFERENCES Ades, S. E. (2008). Regulation by destruction: Design of the σE envelope stress response. Current Opinion in Microbiology, 11(6), 535–540. Akiyama, Y., Kanehara, K., & Ito, K. (2004). RseP (YaeL), an Escherichia coli RIP protease, cleaves transmembrane sequences. The EMBO Journal, 23(22), 4434–4442. Akiyama, K., Mizuno, S., Hizukuri, Y., Mori, H., Nogi, T., & Akiyama, Y. (2015). Roles of the membrane-reentrant beta-hairpin-like loop of RseP protease in selective substrate cleavage. eLife, 4, e08928. Alba, B. M., Leeds, J. A., Onufryk, C., Lu, C. Z., & Gross, C. A. (2002). DegS and YaeL participate sequentially in the cleavage of RseA to activate the σE-dependent extracytoplasmic stress response. Genes & Development, 16(16), 2156–2168. Bai, X. C., Yan, C., Yang, G., Lu, P., Ma, D., Sun, L., et al. (2015). An atomic structure of human γ-secretase. Nature, 525(7568), 212–217. Bolduc, D. M., Montagna, D. R., Gu, Y., Selkoe, D. J., & Wolfe, M. S. (2016). Nicastrin functions to sterically hinder γ-secretase-substrate interactions driven by substrate transmembrane domain. Proceedings of the National Academy of Sciences of the United States of America, 113(5), E509–E518. Brown, M. S., Ye, J., Rawson, R. B., & Goldstein, J. L. (2000). Regulated intramembrane proteolysis: A control mechanism conserved from bacteria to humans. Cell, 100(4), 391–398. Chen, G., & Zhang, X. (2010). New insights into S2P signaling cascades: Regulation, variation, and conservation. Protein Science, 19(11), 2015–2030. Davis, L., & Chin, J. W. (2012). Designer proteins: Applications of genetic code expansion in cell biology. Nature Reviews Molecular Cell Biology, 13(3), 168–182. Dickey, S. W., Baker, R. P., Cho, S., & Urban, S. (2013). Proteolysis inside the membrane is a rate-governed reaction not driven by substrate affinity. Cell, 155(6), 1270–1281. Feng, L., Yan, H., Wu, Z., Yan, N., Wang, Z., Jeffrey, P. D., et al. (2007). Structure of a site-2 protease family intramembrane metalloprotease. Science, 318(5856), 1608–1612. Flynn, J. M., Levchenko, I., Sauer, R. T., & Baker, T. A. (2004). Modulating substrate choice: The SspB adaptor delivers a regulator of the extracytoplasmic-stress response to the AAA+ protease ClpXP for degradation. Genes & Development, 18(18), 2292–2301. Hikita, C., & Mizushima, S. (1992). Effects of total hydrophobicity and length of the hydrophobic domain of a signal peptide on in vitro translocation efficiency. The Journal of Biological Chemistry, 267(7), 4882–4888. Hizukuri, Y., & Akiyama, Y. (2012). PDZ domains of RseP are not essential for sequential cleavage of RseA or stress-induced σE activation in vivo. Molecular Microbiology, 86(5), 1232–1245. Hizukuri, Y., Oda, T., Tabata, S., Tamura-Kawakami, K., Oi, R., Sato, M., et al. (2014). A structure-based model of substrate discrimination by a noncanonical PDZ tandem in the intramembrane-cleaving protease RseP. Structure, 22(2), 326–336. Inaba, K., Suzuki, M., Maegawa, K.-I., Akiyama, S., Ito, K., & Akiyama, Y. (2008). A pair of circularly permutated PDZ domains control RseP, the S2P family intramembrane protease of Escherichia coli. The Journal of Biological Chemistry, 283(50), 35042–35052. Kanehara, K., Akiyama, Y., & Ito, K. (2001). Characterization of the yaeL gene product and its S2P-protease motifs in Escherichia coli. Gene, 281(1–2), 71–79.

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Kanehara, K., Ito, K., & Akiyama, Y. (2002). YaeL (EcfE) activates the σE pathway of stress response through a site-2 cleavage of anti-σE, RseA. Genes & Development, 16(16), 2147–2155. Kanehara, K., Ito, K., & Akiyama, Y. (2003). YaeL proteolysis of RseA is controlled by the PDZ domain of YaeL and a Gln-rich region of RseA. The EMBO Journal, 22(23), 6389–6398. Koide, K., Ito, K., & Akiyama, Y. (2008). Substrate recognition and binding by RseP, an Escherichia coli intramembrane protease. The Journal of Biological Chemistry, 283(15), 9562–9570. Koide, K., Maegawa, S., Ito, K., & Akiyama, Y. (2007). Environment of the active site region of RseP, an Escherichia coli regulated intramembrane proteolysis protease, assessed by sitedirected cysteine alkylation. The Journal of Biological Chemistry, 282(7), 4553–4560. Kroos, L., & Akiyama, Y. (2013). Biochemical and structural insights into intramembrane metalloprotease mechanisms. Biochimica et Biophysica Acta, 1828(12), 2873–2885. Langosch, D., Scharnagl, C., Steiner, H., & Lemberg, M. K. (2015). Understanding intramembrane proteolysis: From protein dynamics to reaction kinetics. Trends in Biochemical Sciences, 40(6), 318–327. Lima, S., Guo, M. S., Chaba, R., Gross, C. A., & Sauer, R. T. (2013). Dual molecular signals mediate the bacterial response to outer-membrane stress. Science, 340(6134), 837–841. Maegawa, S., Koide, K., Ito, K., & Akiyama, Y. (2007). The intramembrane active site of GlpG, an E. coli rhomboid protease, is accessible to water and hydrolyses an extramembrane peptide bond of substrates. Molecular Microbiology, 64(2), 435–447. Melton, D. A., Krieg, P. A., Rebagliati, M. R., Maniatis, T., Zinn, K., & Green, M. R. (1984). Efficient in vitro synthesis of biologically active RNA and RNA hybridization probes from plasmids containing a bacteriophage SP6 promoter. Nucleic Acids Research, 12(18), 7035–7056. Mizushima, S., Tokuda, H., & Matsuyama, S. (1991). In vitro biochemical studies on translocation of presecretory proteins across the cytoplasmic membrane of Escherichia coli. Methods in Cell Biology, 34, 107–146. Moin, S. M., & Urban, S. (2012). Membrane immersion allows rhomboid proteases to achieve specificity by reading transmembrane segment dynamics. eLife, 1, e00173. Rawson, R. B., Zelenski, N. G., Nijhawan, D., Ye, J., Sakai, J., Hasan, M. T., et al. (1997). Complementation cloning of S2P, a gene encoding a putative metalloprotease required for intramembrane cleavage of SREBPs. Molecular Cell, 1(1), 47–57. Saito, A., Hizukuri, Y., Matsuo, E., Chiba, S., Mori, H., Nishimura, O., et al. (2011). Postliberation cleavage of signal peptides is catalyzed by the site-2 protease (S2P) in bacteria. Proceedings of the National Academy of Sciences of the United States of America, 108(33), 13740–13745. Sun, L., Li, X., & Shi, Y. (2016). Structural biology of intramembrane proteases: Mechanistic insights from rhomboid and S2P to γ-secretase. Current Opinion in Structural Biology, 37, 97–107. Urban, S. (2013). Mechanisms and cellular functions of intramembrane proteases. Biochimica et Biophysica Acta, 1828(12), 2797–2800. Wang, Y., Zhang, Y., & Ha, Y. (2006). Crystal structure of a rhomboid family intramembrane protease. Nature, 444(7116), 179–180. Weihofen, A., & Martoglio, B. (2003). Intramembrane-cleaving proteases: Controlled liberation of proteins and bioactive peptides. Trends in Cell Biology, 13(2), 71–78. Wolfe, M. S. (2009). Intramembrane proteolysis. Chemical Reviews, 109(4), 1599–1612. Young, T. S., Ahmad, I., Yin, J. A., & Schultz, P. G. (2010). An enhanced system for unnatural amino acid mutagenesis in E. coli. Journal of Molecular Biology, 395(2), 361–374.

CHAPTER TWO

Signal Peptidase Enzymology and Substrate Specificity Profiling € R.E. Dalbey*,1, D. Pei*, O.D. Ekici† *The Ohio State University, Columbus, OH, United States † The Ohio State University, Newark, OH, United States 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Purification of Δ2-75 SP1 2.1 Reagents 2.2 Methods 3. Purification of SP1 Substrate pONA 3.1 Reagents 3.2 Method 4. In Vitro Signal Peptide Cleavage Assay 4.1 Reagents 4.2 Method 5. In Vivo Assay of Signal Peptidase Activity 5.1 Reagents 5.2 Method 6. Substrate Specificity Profiling 6.1 Peptide Library 6.2 On-Bead Screening of Peptide Library 6.3 Peptide Sequence 7. Conclusions Acknowledgment References

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Abstract Signal peptidases are membrane proteases that play crucial roles in the protein transport pathway of bacteria. They cleave off the signal peptide from precursor proteins that are membrane inserted by the SecYEG or Tat translocons. Signal peptide cleavage releases the translocated protein from the inner membrane allowing the protein to be exported to the periplasm, outer membrane, or secreted into the medium. Signal peptidases are very important proteins to study. They are unique serine proteases with a Ser-Lys dyad, catalyze cleavage at the membrane surface, and are promising potential

Methods in Enzymology, Volume 584 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2016.09.025

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antibacterial drug targets. This chapter will focus on the isolation of signal peptidases and the preprotein substrates, as well as describe a peptide library approach for characterizing the substrate specificity.

1. INTRODUCTION In all cells, proteins exported across membranes are made in a precursor form with an amino-terminal cleavable signal peptide. Signal peptidases, which are unconventional serine proteases, cleave off the signal peptide from the exported proteins to generate the mature protein during or shortly after export. Cleavage is required to release exported proteins from the membrane (Dalbey & Wickner, 1985). Signal peptidases belong to a group of membrane-bound proteases that cleave substrates at the membrane surface (Dalbey, Wang, & van Dijl, 2012). Catalysis occurs on the outside surface of the membrane in a water environment. This is in contrast to intramembrane proteases that cleave their substrates within the membrane plane and possess a water-accessible channel within the membrane (Brown, Ye, Rawson, & Goldstein, 2000; Wolfe & Kopan, 2004). The signal peptidase most thoroughly studied is from Escherichia coli. It was the first signal peptidase to be cloned (Date & Wickner, 1981), sequenced (Wolfe, Wickner, & Goodman, 1983), and purified (Zwizinski & Wickner, 1980); and was also the first to be crystallized (Paetzel et al., 1995) and have its structure solved to high resolution (Paetzel, Dalbey, & Strynadka, 1998). Other well-characterized signal peptidases are the Bacillus subtilis SipS (van Dijl, de Jong, Venema, & Bron, 1995), Streptococcus pneumoniae SP1 (Peng et al., 2001), and the Staphylococcus aureus SpsB (Ting et al., 2016). All bacterial SP1 (signal peptidase) proteases characterized to date possess both a critical serine and a lysine which are needed for catalysis. In 1998, the structure of the catalytic periplasmic domain (Δ2-75) of the E. coli SP1 was solved to 1.9 A˚ in complex with a 5S,6S β-lactam-type inhibitor (Paetzel et al., 1998). The structure showed the active site region with the catalytic Ser-Lys dyad and the potential oxyanion hole residues. Interestingly, the catalytic serine residue formed a covalent adduct with the inhibitor analogous to the acyl intermediate in the proteolytic mechanism, and the lysine residue that functions as a general base is within hydrogen bonding distance of the serine γ-oxygen. Further, the catalytic domain was found to have an exposed hydrophobic surface that interacts with the membrane bilayer, most likely helping SP1 bind and cleave the precursor protein at the membrane surface.

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In addition to the Ser-Lys active site architecture, the structure (Fig. 1) revealed the S1 pocket that binds the
1 residue (also called the P1 residue) of the substrate and the S3 pockets that bind the substrate
3 residue (also

Fig. 1 Signal peptide model with the signal peptide pONA in complex with SP1. (A) The signal peptide depicted in a ribbon diagram is shown in gray. The surface of the Δ2-75 molecule within 5 Å of the signal peptide is shown. The two molecules of phospholipids are depicted to indicate where the membrane would lie. (B) Close-up of the substratebinding region of SP1. The surface of SP1 is shown in green for the residue that is involved in forming the S1 substrate-binding pocket. Cyan shows the molecular surface of those SP1 residues forming the S3 pocket. Isoleucine 86 and isoleucine 144 are shown in red. The
1 and
3 alanine of the pONA signal peptide are shown in yellow, and the SP1 residues are indicated in black. This figure was reproduced with permission from Karla, A., Lively, M. O., Paetzel, M., & Dalbey, R. (2005). The identification of residues that control signal peptidase cleavage fidelity and substrate specificity. The Journal of Biological Chemistry, 280(8), 6731–6741. Retrieved from http://www.ncbi.nlm.nih.gov/entrez/query.fcgi? cmd¼Retrieve&db¼PubMed&dopt¼Citation&list_uids¼15598653.

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called the P3 residue) (Paetzel et al., 1998). The S1 pocket is relatively small allowing it to bind to a small
1 residue of the signal peptide, while the S3 pocket is larger and capable of binding either small or large aliphatic
3 residues of the signal peptide. I144 and I86 of the E. coli enzyme are positioned to contribute to the substrate binding and substrate specificity (Karla, Lively, Paetzel, & Dalbey, 2005). Mutation of I144 to Cys resulted in poor cleavage accuracy at a range of cleavage sites within the substrate proOmpA nuclease A, suggesting that residue I144 is important for the fidelity of cleavage (Karla et al., 2005). Moreover, an I144A/I86A double mutant exhibited a broader specificity as it was capable of cleaving after a phenylalanine at the
1 position of the proOmpA nuclease A (Karla et al., 2005). The wild-type SP1 was not capable of cleaving a preprotein after the
1 Phe. Substrate specificity profiling was used to characterize a variety of E. coli signal peptidase mutants with mutations in the S1 and S3 pockets (Ekici et al., 2007). The profiling was carried out by using a combinatorial peptide library. This library approach is high throughput and can be employed to identify the protease cleavage site. The study showed that the I144A and I144C mutants had an expanded specificity at the
1 and
3 positions (Ekici et al., 2007). Remarkably, a double SP1 mutant (I144C/I86T) cleaved substrates containing a
3 arginine at a similar frequency to those with a
3 alanine, suggesting that the specificity of the mutant enzyme was markedly changed (Ekici et al., 2007). The study thus revealed the critical roles of the I144A and I86 residues in controlling the substrate specificity and cleavage fidelity. Importantly, in 2016, the structure of the S. aureus SP1 (SpsB) was solved with a signal peptide bound using a peptide tethering technique (Ting et al., 2016). The signal peptidase catalytic domain was solved as a fusion protein with the signal peptide of maltose-binding protein bound to the SP1. The structure revealed the S1 and S3 pockets, which are conserved between SpsB and the E. coli SP1. The signal peptide-bound complex reinforced the role of I144 and I86, forming a bridge between the S1 and S3 pocket (Ting et al., 2016). The
2 and
4 residue side chains point away from the protein and are solvent exposed. Ultimately, the S. aureus SpsB structure revealed also the SP1 groove that binds a variety of residues seen in preproteins at the +1, +2, and +3 position. It also showed how the signal peptide was bound to the catalytic domain up to where it would enter the membrane. Signal peptidase is an attractive antibacterial drug target for various reasons (Craney & Romesberg, 2015; Smitha Rao & Anne, 2011). SP1 is conserved

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among bacterial pathogens and has a novel active site architecture (Ser-Lys dyad) that could be selectively targeted, and studies show there are good inhibitors against the SP1. Why do we need new antibiotics? First, antimicrobial resistance to existing antibiotics is growing, causing reduced efficacy. The drug resistance of bacterial pathogens has led to an urgent demand for new treatments (Craney & Romesberg, 2015; Smitha Rao & Anne, 2011). The most promising class of SP1 inhibitors are the arylomycins, which are produced by Streptomyces (Craney & Romesberg, 2015). Romesberg and coworkers have shown that a broad spectrum of bacteria is sensitive to arylomycins. By analyzing the arylomycin-sensitive Staphylococcus epidermidis, they discovered that resistance-causing point mutations occur in the region preceding the catalytic serine residue, and these mutations weaken the binding of SP1 to the arylmycin (Smith, Roberts, & Romesberg, 2010). Analogous mutations in SP1 are also found in a number of bacteria that are resistant to arylomycin compounds (Smith et al., 2010). Romesberg and coworkers attempted to change the arylomycin structure to improve their potency against SP1 (Liu, Smith, Steed, & Romesberg, 2013). SP1s are inhibited by certain β-lactam derivatives, but not by classical protease inhibitors (Black & Bruton, 1998; Craney & Romesberg, 2015; Smitha Rao & Anne, 2011). One β-lactam derivative, allyl (5S,6S)-6-[(R)-acetoxyethyl] penem-3-carboxylate, has an IC50 of 0.38 μM (Black & Bruton, 1998). A natural lipoglycopeptide is quite effective at inhibiting SP1 with an IC50 in the micromolar to nanomolar range (IC50 of 0.11–0.19 μM for E. coli and IC50 of 2.4–24.9 μM for S. pneumoniae (Kulanthaivel et al., 2004)). Its minimum inhibitor concentration is less than 80 μM for these bacteria. This chapter will focus on the enzymology of signal peptidases and substrate profiling methods. We will describe the purification of the E. coli signal peptidase catalytic domain used for solving the structure of the enzyme and also describe the purification of SP1 preprotein substrates. Additionally, we will report a high-throughput peptide library approach to characterize the substrate specificity of SP1 enzymes and proteases in general.

2. PURIFICATION OF Δ2-75 SP1 Here we report the purification of Δ2-75 SP1, the catalytic domain of SP1. We already published the purification of a full-length SP1 with a His tag located internally by changing amino acid residues 35–40 to histidine residues. (Carlos, Paetzel, et al., 2000; Wang & Dalbey, 2010). Δ2-75 SP1, which is missing the two membrane-anchoring segments of SP1, was initially

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constructed by Kuo et al. (1993). Δ2-75 has a hydrophobic surface area that allows it to bind to lipid vesicles and partition into lipid monolayers (van den Brink-van der Laan, Dalbey, Demel, Killian, & de Kruijff, 2001; van Klompenburg et al., 1998). The crystal structure of the Δ2-75 apoenzyme (Paetzel, Dalbey, & Strynadka, 2002) and various SP1 inhibitor complexes has been solved to high resolution (Liu et al., 2011; Luo, Roussel, Dreier, Page, & Paetzel, 2009; Paetzel et al., 1998; Paetzel, Goodall, Kania, Dalbey, & Page, 2004). Below we describe an optimized purification protocol for Δ2-75, which was used to purify the protein for crystallization (Paetzel et al., 1995). This procedure leads to greater than 95% purity of Δ2-75 after washing of the inclusion bodies. Δ2-75 runs as a single band on an SDS-PAGE gel. In the final ion exchange chromatography step (after Δ2-75 has refolded), the nonionic Triton X-100 detergent is removed.

2.1 Reagents Lysis buffer: 20 mM Tris, 5 mM MgCl2, pH 7.4 Solubilizing buffer: 6 M guanidine–HCl, 20 mM Tris–HCl, pH 7.4 Buffer A: 0.5% Triton X-100, 10 mM EDTA, 20 mM Tris–HCl, pH 7.4 Buffer B: 20 mM Tris–HCl pH 7.4

2.2 Methods 1. The E. coli strain BL21 (DE3) is transformed with the pET-3d vector encoding Δ2-75. 2. 1 L of cells are grown at 37°C to an optical density of 0.6 at 600 nm in Luria–Bertani (LB) medium containing 100 μg/mL ampicillin. Δ2-75 is expressed by the addition of 0.5 mM IPTG. Growth of the culture is continued for 4 h. The cells are harvested by centrifugation, resuspended in 8 mL of lysis buffer, and lysed by five passages through a French pressure cell (16,000 psi). 3. The inclusion bodies are isolated by centrifugation (12,000  g, 5 min, 4°C) and washed five times in 10 mL of buffer A. In each step, the solubilized contaminated proteins are separated from inclusion bodies by centrifugation (12,000  g, 5 min, 4°C). 4. Δ2-75 is solubilized in 100 mL of solubilizing buffer at room temperature for 1 h. The solution is slowly added to 200 mL of buffer A and is dialyzed five times in the same buffer. To pellet the aggregates, the solution is centrifuged (15,000  g for 1 h) after dialysis. The supernatant is then loaded

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on a Q Sepharose FF anion exchange column equilibrated in buffer B. The protein is eluted off the column with 0.7 M NaCl in buffer B. This chromatography step removes the Triton X-100 detergent and serves to purify the protein. Following the ion exchange step, dialysis (four times) is used to remove the salt, and the protein is concentrated by ultrafiltration.

3. PURIFICATION OF SP1 SUBSTRATE pONA ProOmpA nuclease A (pONA) is an excellent substrate for SP1 (Chatterjee, Suciu, Dalbey, Kahn, & Inouye, 1995). SP1 processes this substrate very well, more efficiently than peptide substrates. There are several procedures to purify pONA, and they all utilize the same initial steps. These steps include the isolation of pONA in the membrane fraction, solubilization of pONA in the membrane fraction in guanidine/high salt buffer, and the separation of the pONA precursor from the mature form by 55% ammonium sulfate precipitation (only the precursor precipitates). After these steps, different procedures are used for the later steps. In the original procedure (Chatterjee et al., 1995) denaturing sizeexclusion chromatography and SP Sepharose ion exchange chromatography to purify the protein is employed. The second purification procedure (Carlos, Paetzel, et al., 2000; Wang & Dalbey, 2010) takes advantage of a His6-tagged pONA construct and uses nickel chelate affinity chromatography as the final ion exchange chromatography step instead of the SP Sepharose. The third procedure (Carlos, Klenotic, Paetzel, Strynadka, & Dalbey, 2000) is described below in which the denaturing size-exclusion step in Chatterjee et al. (1995) is omitted and the final step was SP Sepharose ion exchange chromatography.

3.1 Reagents TEP buffer: 25 mM Tris–HCl, pH 8.0, 5 mM EDTA, 1 mM PMSF MEB buffer: 50 mM Tris–HCl, pH 8.0, 1 mM EDTA, 1 M KCl, 2 M guanidine–HCl, 3 mM β-mercaptoethanol Dialysis buffer 1: 50 mM Tris–HCl, pH 8.8, 1 M KCl, 10 mM CaCl2, 20% glycerol (w/v) Dialysis buffer 2: 50 mM Tris–HCl, pH 8.8, 1 M KCl, 10 mM CaCl2 Dialysis buffer 3: 50 mM Tris–HCl, pH 8.8, 10 mM CaCl2 Guanidine buffer: 7.2 M guanidine–HCl, pH 3.0, 50 mM sodium citrate, 5 mM β-mercaptoethanol HEPES buffer: 25 mM HEPES, pH 7.5, 1 M NaCl

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3.2 Method 1. The E. coli strain BL21 (DE3) is transformed with the pET-21a bearing the proOmpA nuclease A gene (pONA). 2. 1 L of cells are grown at 37°C to an absorbance of 0.8 at 600 nm in LB medium containing 100 μg/mL ampicillin. pONA is expressed for 3 h by the addition of 0.5 mM IPTG. 3. The cells are centrifuged at 6500  g for 10 min and resuspended in TEP buffer with 5 mL per gram of cells. 1 mM PMSF (final concentration) is added and the cells are French pressed two times at 10,000 psi. 4. The cells lysate is centrifuged (120,000  g) for 1 h, and the membrane pellet containing pONA is resuspended in 20 mL MEB guanidinium buffer. The sample is sonicated three times and stirred (30 min, 4°C). 5. The sample is centrifuged at 230,000  g for 30 min to pellet the membranes. The solubilized pONA in the supernatant is dialyzed overnight at 4°C in dialysis buffer 1. The supernatant is then dialyzed for 4 h in dialysis buffer 2, followed by dialysis for 4 h with dialysis buffer 3. Precipitation of proteins usually occurs in the third dialysis step. Residual aggregates in the dialyzed sample are removed by centrifugation at 230,000  g for 30 min. 6. Ammonium sulfate is added to the sample to 20% saturation (2.3 g of ammonium sulfate per 20 mL) and incubated at 4°C for 30 min. The precipitated proteins that are contaminants are removed by centrifugation at 23,000  g for 30 min. Ammonium sulfate is added to the sample to 55% saturation (4.5 g ammonium sulfate per 20 mL) and incubated at 4°C for 30 min. The precipitated precursor pONA is isolated by centrifugation at 23,000  g for 30 min. The contaminating mature ONA does not precipitate under these conditions. 7. The pONA is resuspended in 5 mL of guanidinium buffer and centrifugation is performed at 230,000  g for 30 min to remove any of the insoluble protein. 8. pONA is slowly refolded at 4°C by dialyzing against 100 mL HEPES buffer, followed by 4 L of HEPES. Centrifugation is performed at 230,000  g for 30 min to remove any nonfolded pONA. 9. Supernatant containing the refolded pONA is loaded onto 15 mL SP Sepharose column equilibrated in HEPES buffer. The pONA is eluted by a 0–1 M NaCl gradient in HEPES buffer and analyzed by 17% SDSPAGE. The cleanest fractions are pooled and the concentration of

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pONA is determined using E1% at 280 nm of 8.3. Typically, small aliquots of the substrate are stored at 108 individual clones. In the obtained library, Nanobodies are displayed as a fusion to a phage coat protein and phage display (biopannings) can be performed to enrich for antigen-specific Nanobodies. Biopanning can be done on different sources of antigen: recombinant protein, whole cells/virus-like particles (VLPs), total membranes, etc., either on solid phase or in solution. To validate the binding of generated Nanobodies to the target protein a variety of assays can be used, e.g., AlphaScreen, In-Cell ELISA, FACS, ELISA, surface plasmon resonance (SPR). Functional assays can provide information on functional effects the Nanobodies possibly exert on the target protein. Further characterization of the selected Nanobodies can include protein-specific (functional) assays, epitope, and affinity determination or other. Depending on the purpose of the generated Nanobodies, they can be used for fundamental biochemical studies, in vivo imaging, protein crystallization, clinical and/or industrial applications.

Yeast (Koide & Koide, 2012; Ryckaert, Pardon, Steyaert, & Callewaert, 2010), bacterial (Fleetwood et al., 2013; Salema et al., 2013; Wendel, Fischer, Martı´nez, Sepp€al€a, & Nørholm, 2016), or ribosome display (Hanes & Pluckthun, 1997; He & Taussig, 2002; Zahnd, Amstutz, & Pl€ uckthun, 2007) are also being used and described elsewhere. More

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recently a novel method using in vitro mRNA/cDNA display has been described by Doshi et al. (2014).

3. CRITICAL ASPECTS OF BIOPANNINGS Nanobodies are isolated from an immune library, which typically contains a few million different clones, by in vitro phage display (biopanning). In each biopanning round, phages interacting specifically with an immobilized antigen are captured, while nonspecific phages are washed away. The phage pool enriched for the antigen is recovered and used to infect Escherichia coli to rescue the phages. A sublibrary enriched for antigen-specific phages is prepared after a biopanning round and may be used in a subsequent round. Alternatively, individual Nanobodies can be expressed and screened in a suitable assay to select Nanobody clones specifically binding to the tested antigen. We monitor the phage enrichment after each biopanning round by performing background selections (i.e., including a parallel selection of phages in identical biopanning conditions, while omitting the antigen of interest) and by comparing the phage titer of both selection conditions (for a method to determine phage titers, see Pardon et al., 2014 step 47). Low enrichments (70%, 3  104 Hi5 insect cells are seeded per well in a 96-well plate to obtain confluent wells after ON incubation.

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(2) Incubate ON in incubator at 27°C. In order to prevent evaporation, create a wet chamber inside the incubator. (3) Discard media and add 100 μL of 4% PFA/1 PBS. (4) Incubate 30 min at RT and then wash 3  with 150 μL 1  PBS. The plates are now ready for use. Alternatively, at this point, 150 μL of PBS can be added and covered plates can be stored at 4°C for maximum a few weeks. For longer storage, a preservative could be added, e.g., 10 mM sodium azide. Protocol: sample loading and plate processing Screening can be performed on intact cells (extracellular binding of Nanobody) or permeabilized cells (intracellular binding of Nanobody). In case the target is extracellular, start at step 3 in the following protocol. (1) Permeabilize the cells with 200 μL of 0.1% Triton in 1  PBS during 1  5 min. (2) Remove 0.1% Triton in 1  PBS. (3) Add 75 μL of periplasmic extracts. (4) Incubate ON at 4°C while gently shaking. (5) Wash 3  with PBS. (6) Add biotinylated anti-HA antibody diluted 1 in 500 in 1  PBS + 2.5% BSA. (7) Incubate 2 h at RT. (8) Wash 3  with 1  PBS. (9) Add streptavidin-HRP as secondary antibody, diluted 1 in 5000 in PBS + 2.5% BSA. (10) Incubate 1 h at RT. (11) Wash 4  with 1  PBS. (12) Add 75 μL of substrate solution. (13) Stop reaction with 75 μL of 2 N H2SO4 when color turns blue. (14) Measure absorbance at 450 nm. Note: Optionally, the amount of cells in each well that were viable before fixation can be measured as control for well-to-well variability. 4.1.3 Sequencing-Based Methods In the absence of a suitable screening assay, picking and sequencing of 100s of clones from each biopanning round is a potential alternative. The number of clones which need to be screened/characterized can be greatly reduced by removing invalid sequences and redundancy (clustering sequences based on homology and selection of only one representative Nanobody per cluster). Nanobodies in the same cluster usually originate from

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the same parental clone, but differ in their sequence due to somatic hypermutation during affinity maturation (or are PCR induced). As they originate from the same parental clone, they most probably bind the same epitope. Alternatively, the rapidly declining cost of high-throughput sequencing (HTS) and gene synthesis opens novel methods to screen for Nanobodies. HTS may be used to sequence the libraries before and after biopanning. Sequences that are enriched after biopanning are more likely to bind the target protein. One drawback is, as sequencing happens on the whole library and not on single colonies, that sequences are only available in silico. Hence, the DNA has to be synthesized in order to further characterize the selected clones. Examples of this method can be found in Fridy et al. (2014) and Miyazaki et al. (2015).

4.2 Functional Screenings 4.2.1 AlphaScreen Competition AlphaScreen can be used to screen for Nanobodies that disrupt a specific interaction (e.g., receptor/ligand, antibody/antigen, etc.). In this case, each of the interacting proteins is bound to acceptor or donor beads, the pair-wise interaction brings the beads into proximity, and the signal is generated. Nanobodies disrupting this interaction will be identified as hits. Several examples of competition assays using AlphaScreen technology have been described by Rouleau, Turcotte, Mondou, Roby, and Boss
e (2003) and Taouji, Dahan, Bosse, and Chevet (2009). 4.2.2 Screening for Nanobodies Regulating γ-Secretase Function Screenings of Nanobodies can also be performed using functional in vitro assays. However, functional assays are not always compatible with crude periplasmic extract. Furthermore, the concentration of the Nanobody in periplasmic extract may not be sufficiently high to elicit a clear functional effect. Therefore, enrichment of Nanobodies from periplasmic extract and removal of periplasmic extract matrix can be crucial to design a robust functional-based screening assay. In order to do this, the antigen of interest can be immobilized on a solid carrier (e.g., a 96-well plate coated with NHS functional groups (as described below)), followed by addition of periplasmic extracts containing Nanobodies. While antigen-specific Nanobodies will be captured by the antigen, other nonrelevant proteins will be washed away. We estimate that Nanobodies interacting with relatively high affinity could reach concentrations around 1 μM or higher in the assay, in a total volume of 40 μL per well.

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We have determined the optimal amount of γ-secretase to be coupled to the plate (
20 nM final in activity assay) to obtain a functional readout (Fig. 4). This concentration is preferably 10- to 100-fold lower than the estimated concentration of Nanobody in the assay, to be able to detect an effect of the Nanobody. For γ-secretase, we then carry out an in vitro activity assay with γ-secretase substrate and measure Aβ production by ELISA (Cha´vezGuti
errez et al., 2008). Materials (1) Purified γ-secretase (2) 3D-NHS 96-well plate, white, flat bottom (PolyAn, catalog number 00680451) (3) Coupling buffer (0.2 M NaHCO3, 0.5 M NaCl pH 8.3, 0.25% CHAPSO)

Fig. 4 In vitro γ-secretase activity assays for the screening of γ-secretase modulating Nanobodies. γ-Secretase activity assay on solid support used as functional screening for the selection of Nanobodies regulating protease function. Upper panel: Different concentrations of the protease were coated on NHS plates and then activity assays were performed and Aβ 40 production quantified by ELISA. Lower panel: Periplasmic extracts containing anti-γ-secretase Nanobody 28 (anti-Nicastrin) and Nanobody 30 (complexspecific) were incubated with immobilized purified γ-secretase or immobilized purified GFP (control) for 12 h. Wells were washed and proteins captured were eluted. Elution fractions loaded on western blot and stained with anti-His antibody show enrichment of Nanobodies on the protease complex.

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(4) Blocking buffer (100 mM Tris–HCl pH 8.5, 0.25% CHAPSO) (5) 100 μL periplasmic extracts of Nanobodies to screen (6) Washing buffer (25 mM PIPES pH 7.4, 150 mM NaCl, 0.1% CHAPSO) Protocol (1) Add purified γ-secretase to the well to a final concentration of 20 nM in 40 μL coupling buffer. (2) Incubate ON at 4°C while gently shaking at 200 rpm. (3) Collect unbound fractions and measure protein concentration. Input vs unbound sample concentrations inform about coupling efficiency. (4) Block plate with 100 μL of blocking buffer during 30 min. Repeat this step 2 times more, with incubation steps of 30 min and 1 h, respectively. (5) Add 100 μL periplasmic extracts containing Nanobodies and incubate ON at 4°C. (6) Remove unbound fractions and wash plate 3  with 100 μL of washing buffer. (7) Perform functional assay. For γ-secretase, an in vitro activity assay as described in Cha´vez-Guti
errez et al. (2008) is carried out. Detection of Aβ 40/42 products is performed by ELISA.

5. CHARACTERIZATION OF NANOBODIES After the screening assay(s), we sequence and cluster the positive clones according to sequence homology, as mentioned in Pardon et al. (2014). One Nanobody for each family is selected and Nanobodies are expressed and purified for further characterization. Most screening assays can be reused for characterization, but now dilution series of the selected, purified Nanobodies are tested in order to compare their respective potencies/affinities in these assays. Here we list a limited set of potential characterization assays.

5.1 Defining the Epitope Characterization of the epitope may provide valuable insights into the structural/functional relationships of the protein of interest. Furthermore, this information can be used to determine which Nanobodies are compatible to set up a sandwich ELISA or to generate multivalent Nanobodies (see also Section 6). Antibody (fragments) targeting the same epitope frequently shares the same functional effect (Klein et al., 2013; Markovitz, Healey,

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Parker, Meeks, & Lollar, 2013). Conversely, if Nanobodies from multiple epitope bins exhibit functional activity, this may imply different mechanisms of action, which can be advantageous if more than one biological pathway needs to be targeted (Emde et al., 2011; Jamnani et al., 2012; Koefoed et al., 2011; Robak et al., 2012; Spangler, Manzari, Rosalia, Chen, & Wittrup, 2012). Delineating the actual epitope of Nanobodies is challenging as many of them are conformation-sensitive probes. However, grouping Nanobodies in clusters that bind the same epitopes (epitope binning) provides in most cases enough information for further experimental setup. 5.1.1 Epitope Binning One method to conduct epitope binning is surface plasmon resonance (SPR). In this setup, the antigen is coupled to the chip and a first Nanobody binds until saturation is reached and then a second Nanobody is injected. If additional binding is observed, Nanobodies belong to a different epitope bin (Fig. 5A). The advantage of this method is that assays can be performed with periplasmic extracts. However, analysis of multimeric membrane protein complexes (such as γ-secretase) is challenging. A few examples of the use of SPR for epitope binning can be found in Hmila et al. (2010) and Steeland et al. (2015). Alternative to SPR, biolayer interferometry (BLI) or Meso Scale Discovery technology can be used (Abdiche et al., 2014; Estep et al., 2013). An alternative binning method is competitive phage ELISA (De Tavernier et al., 2016). In this assay, the antigen captured on a solid surface interacts with a Nanobody clone (in periplasmic extract); then phages displaying a second Nanobody are added. After several washing steps, an HRP-conjugated antiphage antibody (e.g., anti-M13 antibody) reports on the binding of the second Nanobody. Phage binding is only observed if the Nanobody displayed on the phage targets a different epitope than the Nanobody. Be aware that, when the Nanobody displayed on the phage has a much higher affinity than the Nanobody in the periplasmic extract, additional binding might be observed, even if both Nanobodies bind the same epitope, causing a false positive result. To bin our anti-γ-secretase Nanobodies, immunoprecipitation assays followed by SDS-PAGE/western blot analyses are carried out (Fig. 5B). In this assay, a Nanobody covalently coupled to NHS beads interacts with detergent-solubilized γ-secretase in the presence or absence of a second, noncoupled Nanobody. Efficient immunoprecipitation of γ-secretase indicates that the tested Nanobodies bind different epitopes.

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A

B

Response units

1 and 2 in different epitope bins

Start injection Nanobody 1

1 and 2 in same epitope bin Start injection Nanobody 2 Time

C

D

Fig. 5 Characterization of anti-γ-secretase Nanobodies. (A) Theoretical model of epitope binning by SPR. When two Nanobodies belong to different epitope bins, additional binding will be observed. (B) Left panel: Overview of competition experiment. Right panel: Total lysates of insect cells overexpressing human γ-secretase were preincubated with the purified Nanobody (1 μM final concentration) indicated in green. The mix was then tested in pull-down experiments, using immobilized purified Nanobody (1 mg/mL beads) indicated in blue. Western blot against human γ-secretase subunits of pulleddown fractions indicates that Nanobodies 21 and 30 bind the same epitope, while Nanobody 42 belongs to a different bin. (C) Left panel: 250 ng of BSA and purified γ-secretase were dotted on a nitrocellulose membrane and incubated with purified Nanobody (5 μg/mL) or no Nanobody as control. Nanobody binding was detected using anti-HA-antibody (1/4000, Covance, catalog number MMS-101P) and HRP-conjugated anti-mouse antibody (1/10,000, Biorad, catalog number 1721011). Right panel: Western blot against purified human γ-secretase, stained with anti-Nicastrin Nanobody or no Nanobody (control). The arrow indicates the expected MW of Nicastrin. (D) AlphaScreen assay to determine affinity of anti-γ-secretase Nanobody 30. Purified biotinylated human γ-secretase complex (PSEN1/APH1a) and purified Nanobody 30-6XHis were bound by donor and acceptor beads, respectively.

5.1.1.1 Competition Experiment

Materials (1) Purified Nanobodies in solution (2) Purified Nanobodies coupled to NHS beads (see Section 3.2.3.2) (3) Total lysates of cells overexpressing γ-secretase or protein of interest

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(4) Washing buffer: 25 mM PIPES pH 7.4, 150 mM NaCl, 0.5% CHAPSO (5) Elution buffer: 2  lithium dodecyl sulfate (LDS) sample buffer (ThermoFisher Scientific, catalog number NP0008) containing 2% β-mercaptoethanol Protocol (1) Incubate 100 μg of total lysates of cells overexpressing γ-secretase with 1 μM of purified Nanobody for 2 h. (2) Add Nanobodies coupled to NHS beads to total lysates–Nanobody mixes and incubate ON at 4°C while rotating the sample gently head over head. As control for immunoprecipitation, add coupled Nanobodies to total lysates that were not preincubated with Nanobodies. As control for competition, add total lysates to coupled Nanobodies that were preincubated with the same Nanobody. (3) Collect unbound fractions after spinning down beads at 2000  g during 2 min. (4) Wash beads 3 with washing buffer. (5) Elute bound fractions with 2  sample buffer containing 2% β-mercaptoethanol and load on SDS-PAGE/western blot. Stain membrane for protein of interest. We typically use anti-γ-secretase antibodies to detect individual γ-secretase subunits.

5.1.2 Epitope Delineation Different methods can be applied in the search for the epitope. The most precise method relies on the elucidation of the structure of the Nanobody– antigen complex (by NMR, X-ray crystallography, cryo-EM, etc.) (e.g., Du et al., 2013; Eylenstein et al., 2016; Sˇkerlova´ et al., 2015). However, obtaining a structure is not always straightforward. Nonstructure-based methods are described in the literature (Westwood & Hay, 2001), and their applicability depends on the type of epitope: linear or conformational. Most Nanobodies recognize conformational epitopes. Because of their protruding CDR3 region, they preferentially bind conformational epitopes in cavities or clefts (Muyldermans, 2013). Nevertheless, several Nanobodies that bind linear epitopes have been identified before (De Genst et al., 2010; Habib et al., 2013; Itoh et al., 2014; Smolarek et al., 2010). An easy method to check if a Nanobody targets a linear or a conformational epitope is blotting (Fig. 5C). Purified antigen is analyzed by western blot under native (native gels or dot blot) or denaturing conditions

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(SDS-PAGE). Conformational Nanobodies will bind the protein under native conditions, but the interaction with the epitope will not take place upon denaturation of the antigen. Nanobodies binding the antigen on western blot most likely recognize a linear epitope. However, proteins can (partially) refold after or during transfer to a membrane, which would enable the Nanobody to bind a conformational epitope on western blot (Zhou, Chen, Purcell, & Emerson, 2007). Delineation of the epitope can be performed by classical alanine scanning studies and detection of binding to mutated antigens (Vercruysse et al., 2013) or binding to truncated variants of the protein (Zhou et al., 2011). For linear epitopes, phage peptide display libraries can be used, in combination with multiple sequence alignment (Ganglberger et al., 2000; Malik et al., 2016; Thueng-in et al., 2012). Delineation of a conformational epitope is more challenging. Mutation studies cannot always be applied because the mutation itself may cause conformational changes in the protein and therefore interfere with Nanobody binding. One approach is shotgun mutagenesis, which is based on large-scale mutagenesis combined with testing the native folding of proteins in a cellular based assay. This technique has been successfully applied to identify epitopes of GPCR-binding Nanobodies (Davidson & Doranz, 2014; Paes et al., 2009). Another option is to cross-link the Nanobodies to their target, followed by sequencing of the cross-linked target peptide via mass spectrometry as described by Pleiner et al. (2015). Alternatively, a peptide library could be screened to identify the peptides that mimic the conformational epitope on the antigen (also known as mimotopes). Sequences that are enriched after several biopanning rounds could have residues in common with the conformational epitope on the antigen. Further bioinformatic analysis to map these residues on the antigen can be performed using several online tools (a few examples are EpiSearch, Mapitope, and Findmap; an overview can be found in Sun et al., 2013). For membrane proteins, it can be of interest to know if the Nanobodies bind intracellularly or extracellularly. To determine this, the in-cell ELISA described earlier can be used.

5.2 Affinity Determination Nanobody affinity typically lies in the ranges of 105 to 106 M 1 s1 for kon and 102 to 104 for koff rate constants, implying that equilibrium

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dissociation constants in the nano- to picomolar range can be obtained (Muyldermans, 2013). Different methods can be applied in order to determine Nanobody affinity, for example, SPR or BLI. AlphaScreen and ELISA can be performed if these techniques are not available in the lab or if the investigated antigen is not compatible with those assays. We have applied AlphaScreen (similar assay setup as described earlier in Section 4.1.1.2, but now with serial dilutions of purified Nanobody) (Fig. 5D). The assay is carried out at equilibrium and the Nanobodies are titrated while keeping the concentration of the target protein constant. Finally, for membrane proteins, the previously mentioned FACS assay can be used to assess relative affinities, again by titrating the purified Nanobodies against antigenexpressing cells.

6. POTENTIAL APPLICATIONS Nanobodies have been shown to have a broad application range varying from basic fundamental research to therapeutic applications in humans. In the next section, we will briefly discuss diverse Nanobody applications that have recently been developed.

6.1 Fundamental Biochemical and Cellular Studies Nanobodies that bind linear epitopes can be used in fundamental research to detect proteins in immunohistochemistry and western blot (Iqbal et al., 2010; Rakovich et al., 2014; Zheng et al., 2014), although they are not often identified. Antigen-specific Nanobodies can be generated and incorporated in the purification process of (membrane) proteins. Some examples of protein purifications where Nanobodies are used can be found in Pleiner et al. (2015) and are also reviewed in De Marco (2011). In Elad et al. (2015), a conformational Nanobody was used to obtain highly pure and active γ-secretase that could be used for structural analysis. Besides their use as detection tools in vitro, they can also be expressed inside cells as intrabodies for the detection of the target protein. The fact that intrabodies display high specificity for their target at endogenous expression levels of the protein of interest makes them interesting tools to study protein function directly in its native cellular environment. In contrast to conventional antibodies, Nanobodies can readily be expressed

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in the reducing cytoplasmic environment (Staus et al., 2014; Vercruysse, Pardon, Vanstreels, Steyaert, & Daelemans, 2010). Depending on the presence and nature of the signal peptide, they can be directed to several cellular compartments, like the nucleus, mitochondria, ER, or cytoplasma. Protein localization of the target protein is usually assessed by chromobodies (fluorescently tagged Nanobodies) (Kaiser, Maier, Traenkle, Emele, & Rothbauer, 2014). Several articles extensively describe the use of intrabodies for protein knockdown (Marschall, D€ ubel, & B€ oldicke, 2015).

6.2 Crystallization of Proteins Nanobodies have also been found to be able to stabilize highly flexible proteins, including membrane proteins, in specific conformations, thereby reducing conformational heterogeneity and increasing the probability of obtaining a crystal structure. This has been described for BACE2, the β-2 adrenergic receptor, and several other proteins (Banner et al., 2013; Chaikuad et al., 2014; Domanska et al., 2011; Rasmussen, Choi, et al., 2011; Rasmussen, DeVree, et al., 2011). Furthermore. Nanobodies increase the hydrophilic surface area available to form crystal contacts (for a review, see Lieberman, Culver, Entzminger, Pai, & Maynard, 2011). Therefore, Nanobodies are often used as “chaperones” for crystallization of proteins.

6.3 Potential Clinical and Industrial Applications Much research has been focused on the development of Nanobodies as modulators of protein function or as blocking agents of protein–protein interactions (e.g., receptor–ligand interactions), not only to answer fundamental scientific questions but also largely because of potential therapeutic applications. Functional Nanobodies have been generated against a wide variety of antigens, including challenging targets like GPCRs (reviewed in Cromie, Van Heeke, & Boutton, 2015; Muji
c-Deli
c, De Wit, Verkaar, & Smit, 2014) or ion-gated, ligand-gated, and voltage-gated ion channels (L€ ow et al., 2013; www.ablynx.com). Another important feature is their enhanced stability, which opens opportunities for alternative delivery routes. The Belgian biotech company Ablynx has developed a trivalent Nanobody against respiratory syncytial virus (RSV), a virus causing lower respiratory tract infections primarily in infants, which can be administered by nebulization (Detalle et al., 2016). Nebulization requires robust and highly stable biologics. In addition, several

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groups have reported promising results for potential oral delivery of Nanobodies. For example, Harmsen, Van Solt, Van Zijderveld-Van Bemmel, Niewold, and Van Zijderveld (2006) selected Nanobodies with enhanced stability in gastric and jejunal fluids allowing to administer therapeutically relevant doses orally. Other groups have reported effective oral delivery of Nanobody expressing Lactobacilli in various disease settings (e.g., rotavirus-induced diarrhea (Pant et al., 2006), chronic colitis (Vandenbroucke et al., 2010), or Clostridium difficile infections (Andersen et al., 2015)). As Nanobodies are single-chain antibodies, they can easily be cloned in multivalent formats (up to heptavalent (www.ablynx.com)), which allows to hit multiple epitopes or targets with one molecule or to increase avidity. For example, multispecific Nanobodies bind different targets (e.g., clinical candidates ALX-0761 (IL-17A/IL-17F/human serum albumin) and ALX-0061 (IL-6R/human serum albumin)), multivalent Nanobodies can bind identical targets (e.g., Detalle et al., 2016; Huet et al., 2014; Ulrichts et al., 2011), while bi-paratopic Nanobodies bind different epitopes on the same target (De Tavernier et al., 2016; Hufton et al., 2014; Hultberg et al., 2011; J€ahnichen et al., 2010; Moayeri et al., 2015). The different Nanobody building blocks are usually linked together with a flexible glycine–serine linker with variable length. The increased valency of the Nanobodies often leads to an increased overall potency. Currently Ablynx has eight Nanobodies in clinical development and the launch for their first potential product (caplacizumab) for the treatment of acquired thrombotic thrombocytopenic purpura is expected in 2018 (Detalle et al., 2016; Peyvandi et al., 2016; www.ablynx.com). Besides applications in a biologic/therapeutic area, researchers have exploited the Nanobodies’ superior resilience to temperature, pH, and organic solvents for the analysis of environmental chemicals (e.g., caffeine, toxins, and methotrexate), reviewed in Bever et al. (2016).

6.4 Nanobodies for In Vivo Imaging Nanobodies are rapidly gaining importance as noninvasive diagnostic tools (in vivo imaging). Besides their assumed low immunogenicity (due to their high sequence identity to the human VH) (Hassanzadeh-Ghassabeh, Devoogdt, De Pauw, Vincke, & Muyldermans, 2013) and cheaper production cost (Harmsen & De Haard, 2007), their small size is of special interest. Moreover, because of their small size and short in vivo half-life (minutes,

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compared to days for a conventional antibody), they show superior targetto-background signals and allow faster imaging compared to classical mAbs (Huang et al., 2008; Keyaerts et al., 2015; Lemaire et al., 2014; Oliveira et al., 2012; Oliveira, Heukers, Sornkom, Kok, & van Bergen En Henegouwen, 2013). Faster imaging allows incorporation of short-lived radioisotopes, which seriously reduces the exposure of patients to radiation. In addition, their small size allows superior tissue penetration (HassanzadehGhassabeh et al., 2013). However, a downside of their smaller size is their rapid clearance by the kidney, which can result in high doses of labeled Nanobody that accumulates in the kidney and bladder, making detection of the tracer in the vicinity of these organs more difficult and might induce unwanted side effects. Nonetheless, several strategies have been pursued successfully to lower kidney accumulation with promising results (D’Huyvetter, Vincke, et al., 2014; Tchouate Gainkam et al., 2011). For imaging, most reported applications using Nanobodies have employed positron emission tomography (PET) or single-photon emission computed tomography (SPECT), although optical and ultrasound-based imaging are also emerging (for an excellent review, see Chakravarty, Goel, & Cai, 2014). More recently, Nanobodies gained interest in the field of targeted radionucleotide therapy in cancer, largely with the same (dis)advantages as described here for Nanobodies as diagnostic tools (reviewed in D’Huyvetter, Xavier, et al., 2014). Alternatively, targeted delivery of toxins has recently been described by Li et al. with promising preliminary results (Li et al., 2016).

ACKNOWLEDGMENTS This work is supported by a European Research Council grant (ERC), the Fonds voor Wetenschappelijk Onderzoek (FWO), Agentschap Innoveren & Ondernemen, Interuniversity attraction pool IAP P7-16, KU Leuven, VIB, Vlaams Initiatief voor Netwerken voor Dementie Onderzoek (VIND, Strategic Basic Research Grant 135043), Cure Alzheimer’s Fund, and a Methusalem grant from KU Leuven and the Flemish Government. B.D.S. is the Bax-Vanluffelen Chair for Alzheimer’s Disease and is supported directly by the Opening the Future campaign of the Leuven Universiteit Fonds (LUF).

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CHAPTER FOUR

Probing the Activity of Eukaryotic Rhomboid Proteases In Vitro B. Cordier, M.K. Lemberg1 Zentrum f€ ur Molekulare Biologie der Universit€at Heidelberg (ZMBH), DKFZ-ZMBH Allianz, Heidelberg, Germany 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Expression and Solubilization of Rhomboid Proteases From Bacteria 2.1 Parameters to Adjust 2.2 Expression and Membrane Preparation 2.3 Solubilization 2.4 Purification 3. In Vitro Probing of Rhomboid Protease Activity 3.1 Serine Hydrolase Probe to Test Reactivity of Active Site Residues 3.2 EnzChek Protease Assay for Efficient Screening of Rhomboid Protease Activity 3.3 Rapid Testing of In Vitro-Generated TM Domain Substrates 3.4 Cleavage of Chimeric or Tagged Substrates Acknowledgments References

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Abstract Proteolysis within the membrane is a recent concept in biology. Rhomboid intramembrane serine proteases are conserved in evolution and serve as key switches in diverse cellular pathways ranging from signaling to protein degradation. Since deregulation of intramembrane proteolysis can lead to severe diseases including neurodegenerative disorders, dissecting their enzymatic function and specificity becomes crucial. As membrane proteins, their solubilization, and purification are technically challenging. As a start point for a comprehensive in vitro characterization of eukaryotic rhomboid proteases, we depict in this chapter a robust workflow to find the best conditions to obtain pure and active enzymes from a bacterial expression system. To monitor the integrity of their active site and visualize substrate cleavage, various established activity assays including activity-based labeling and gel-based cleavage assays are described. These methods are illustrated by use of the Escherichia coli rhomboid protease GlpG and human RHBDL2 as an example.

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1. INTRODUCTION In recent years, several independent findings pointed toward the existence of unusual proteases that cleave their substrates in transmembrane (TM) domains. As a hydrolytic process, cleavage of peptide bonds in the plan of the membrane was difficult to anticipate; existence of regulated intramembrane proteolysis was only slowly accepted in the protease field. Since the first high-resolution structures of intramembrane proteases revealed the architecture of active sites, the concept of intramembrane cleavage became fully accepted. So far, four different families of intramembrane proteases are known: site-2 metalloproteases, GxGD aspartyl proteases, Rce1-type glutamyl proteases, and rhomboid serine proteases (Lemberg, 2013; Strisovsky, 2016; Sun, Li, & Shi, 2016; Vinothkumar & Freeman, 2013). A unifying theme of these enzymes is that the aqueous active site is formed within the core of a polytopic membrane protein. Within the membrane core, the active site residues or prosthetic zinc atoms are being buried. Despite recent progress, key mechanistic questions remain unsolved. In order to dissect intramembrane proteases reaction cycles, defined in vitro assays are needed. Rhomboids, which are investigated in this chapter, consist of a six-TM domain core with a serine and histidine forming a conserved catalytic center (Lemberg et al., 2005; Wang, Zhang, & Ha, 2006). Whereas a number of recent biochemical studies provided key insights into the enzymology of bacterial rhomboid proteases (Arutyunova et al., 2014; Baker, Young, Feng, Shi, & Urban, 2007; Cho, Dickey, & Urban, 2016; Dickey, Baker, Cho, & Urban, 2013; Lemberg et al., 2005; Moin & Urban, 2012; Schafer, Truong, Otzen, Lindorff-Larsen, & Wolynes, 2016; Strisovsky, Sharpe, & Freeman, 2009; Urban & Wolfe, 2005; Xue & Ha, 2012; Zoll et al., 2014), the more challenging expression and purification of active eukaryotic rhomboid proteases hamper their analysis. Since several eukaryotic rhomboid proteases have additional domains fused to the conserved sixTM-domain core (Lemberg & Freeman, 2007), their detailed functional characterization may reveal new regulatory principles. For instance, a recent biochemical characterization of the Drosophila Rhomboid-4 revealed a calcium-mediated substrate-gating mechanism (Baker & Urban, 2015). In this chapter, we describe robust procedures for the expression and solubilization of rhomboid proteases with the focus on rapid screening procedures including activity-based profiling (Lazareno-Saez, Arutyunova,

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Coquelle, & Lemieux, 2013; Sherratt, Blais, Ghasriani, Pezacki, & Goto, 2012; Vosyka et al., 2013; Wolf, Seybold, Hadravova´, Strisovsky, & Verhelst, 2015; Xue & Ha, 2012) that allows efficient optimization of conditions. To showcase the workflow, we chose the bacterial rhomboid protease GlpG, which is currently the best in vitro studied rhomboid protease (Guo et al., 2016; Lemberg et al., 2005; Paslawski et al., 2015; Urban & Wolfe, 2005; Wang et al., 2006; Zoll et al., 2014), and the human rhomboid protease RHBDL2 that previously has been successfully purified (Lemberg et al., 2005).

2. EXPRESSION AND SOLUBILIZATION OF RHOMBOID PROTEASES FROM BACTERIA 2.1 Parameters to Adjust Over the past few decades, the interest concerning the study of membrane proteins is growing, and their role in various cellular processes started to be unveiled. Nevertheless, owing to their hydrophobicity, the in vitro study of membrane proteins can be highly challenging: among the more than a hundred thousand X-ray resolved structures in the Protein Data Bank (PDB) (http://www.rcsb.org/pdb/home/home.do), only 3% represent membrane proteins. Common biochemical techniques are mainly optimized for watersoluble proteins. Thus, approaches have to be adjusted to the special needs of membrane proteins. The challenge to produce and purify them is susceptible to many factors: membrane proteins are naturally found at low abundance in the cell and the overexpression of recombinant membrane proteins is often toxic for the cell. This toxicity can be explained by the saturation of biogenesis and translocation machineries. To reduce or overcome this problem, a wide range of approaches is available. In order to produce a eukaryotic or a prokaryotic membrane protein, different overexpression systems are used. They all have their own advantages and drawbacks. The system has to be chosen according to the requirements of the protein of interest (e.g., posttranslational modifications) and practical considerations. As a nonexhaustive list, yeast cells, baculovirus insect cell system, mammalian cells, or cell-free systems can be considered (Geng, Bush, Mosyak, Wang, & Fan, 2013; Jidenko et al., 2005; Lu et al., 2014; Schwarz et al., 2007). More exotic systems were recently used to efficiently produce membrane proteins such as the eye of transgenic Drosophila melanogaster (Hackmann, Joedicke, Panneels, & Sinning, 2015).

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However, bacteria, and especially E. coli, are probably the most widely used overexpression systems because of their low cost and their high yield. Beyond the choice of an adequate expression organism, many other parameters have to be screened and adjusted if bacteria are favored. These include the bacterial strain and expression plasmid, growth conditions, buffers, lysis method, and solubilization conditions. Several reviews and method articles were written in the last years to list and detail these screening procedures for expressing membrane proteins (Roy, 2015) and even rhomboid proteases in bacteria (Panwar & Lemieux, 2014). For this chapter, we suggest a general workflow to express and purify the bacterial model rhomboid protease GlpG from E. coli and showcase optimization for the eukaryotic rhomboid protease RHBDL2.

2.2 Expression and Membrane Preparation Described later is a general method to prepare the membrane fraction of E. coli cells expressing rhomboid proteases of interest for subsequent solubilization and purification (Fig. 1). First, the gene encoding the rhomboid is cloned into the bacterial expression vector pET-25b(+) (Fig. 1A). The pET system is widely used for expression of recombinant proteins in E. coli, which is under control of the strong T7 bacteriophage promoter. When transformed into a BL21 (DE3)-pLysS strain (or derivatives) carrying a chromosomal copy of the T7 RNA polymerase, expression is induced by the addition of isopropyl β-D-1-thiogalactopyranoside (IPTG) to the culture medium. The pET-25b(+) plasmid encodes a hexa-histidine tag (His6) which is fused to the C-terminus of the protein. Next, the construct expressing glpG is transformed into the E. coli BL21 (DE3)-pLysS strain. The plasmid encoding rhbdl2 is transformed into the Rosetta™ 2 strain. This strain is designed to express heterologous proteins overcoming codon bias as it carries a plasmid with tRNA genes decoding codons rarely used in E. coli. As it is impossible to predict the behavior of an overexpressed membrane protein and its toxicity for the cell, other plasmid and bacterial strains pairs can be considered. BL21 derivative strains such as C41/C43 (DE3) and more recently Lemo21(DE3), also known as “Walker” strains, are better suited for overexpression of membrane proteins (Miroux & Walker, 1996; Wagner et al., 2008). Some overexpressed proteins can misfold and accumulate in inclusion bodies. Therefore, the use of an expression plasmid containing an araBAD promoter (inducible with arabinose), which is

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Fig. 1 Rhomboid expression and purification workflow using E. coli cells. (A) E. coli glpg and human rhbdl2 genes are cloned into pET25b(+) expression vector. Encoded proteins are fused with a His6tag at the C-terminus. (B) The entire procedure, from bacterial transformation to protein purification, covers a 4-day period depending on the duration of expression. Expression of GlpG and RHBDL2 is done for 16 h at 16°C to help protein folding and reduce the formation of inclusion bodies. Intermediate steps are not depicted.

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weaker than the T7 promotor, was fruitful in the study of the Haemophilus influenza rhomboid protease GlpG (Lemieux, Fischer, Cherney, Bateman, & James, 2007). 2.2.1 Duration The whole procedure, starting from bacterial transformation up to the affinity purification step, can last for 4 or 5 days depending on the duration of protein expression (Fig. 1B). 2.2.2 Materials and Equipment – E. coli strains: Rosetta™ 2 (Novagen) or BL21 (DE3)-pLysS (Novagen) (carrying a chloramphenicol resistance marker) – Expression plasmids pET25b(+) (Novagen) (carrying an ampicillin resistance marker) – LB media – LB agar plates (supplemented with suitable antibiotics) – Ampicillin – Chloramphenicol – 2
Laemmli sample buffer (supplemented by 5% β-mercaptoethanol) (2XSB + BME) – IPTG (stock solution 1 M) – Lysozyme (stock solution 20 mg/mL) – Phenylmethylsulfonyl fluoride (PMSF) (stock solution 100 mM) – Benzonase – n-Dodecyl-β-D-maltoside (DDM) (Anatrace, USA) – Triton X100 (T-X100) (Applichem, Germany) – Nonidet P-40 (NP-40) (Applichem, Germany) – Cholesteryl hemisuccinate Tris salt (CHS) (Anatrace, USA) – Protein concentration estimation: bicinchoninic acid (BCA) assay (ThermoFisher, USA) or Bradford protein assay (Applichem, Germany) kits – Coomassie blue staining solution (Coomassie Brilliant Blue R-250) – Penta-his antibody (QIAGEN, Germany) – Incubator shaker (with adjustable temperature, from 16 to 37°C) – Erlenmeyer cell culture flasks (5 L) – Microcentrifuge tubes – Table top centrifuge – Spectrophotometer – Water bath/Eppendorf ThermoMixer®

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Refrigerated centrifuge 1-L centrifuge bottles 50-mL tubes 15-mL tubes Instrument for lysis (high pressure homogenizer Emulsiflex/French press) Homogenizing pestle Ultracentrifuge tubes Ultracentrifuge SDS-PAGE migration systems and blot system PVDF membrane Imager ImageQuant LAS 4000 (GE Healthcare, United Kingdom)

2.2.3 Methods Transformation • Thaw competent BL21 (DE3)-pLysS or Rosetta™ 2 E. coli cells on ice (the competent cells can be prepared in advance using the CaCl2 method (Sambrook, Fritsch, & Maniatis, 1989). • Add 50 μL of competent E. coli cells into a sterile 1.5-mL microcentrifuge tube. • Add 1–2 μL of expression plasmid (50 ng/μL) to the competent cells and mix (a negative control without expression plasmid can be done in parallel). • Incubate the tube on ice for 30 min. • Incubate the tube in a water bath at 42°C for 1 min. • Incubate the tube on ice for 2 min. • Incubate the tube at room temperature for 2 min. • Add 450 μL of LB medium and incubate for 1 h at 37°C in a water bath or thermomixer. • Spread 150 μL of transformed cells on LB agar plates containing ampicillin (100 μL/mL—Amp100) and chloramphenicol (34 μL/mL— Cm34). Grow overnight at 37°C. Cell culture and protein expression (for a 3-L culture) • Precultures: inoculate freshly transformed colonies into a 1-L flask containing 100 mL LB Amp100 Cm34 (1:1000 dilution for each antibiotic). • Incubate at 37°C in an orbital shaker till the cultures reach an OD600 of approximately 0.6 (3–4 h of growth). • Place three 5-L flasks containing 1 L of LB Amp100, Cm34 (each) on a shaker platform in 30°C incubator. Shake to warm and aerate.

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Large-scale cultures: inoculate the cultures at OD600 0.02 (prewarmed and preaerated). • Grow at 30°C to OD600 0.3/0.4. • Remove a 1-mL sample from the culture (cells not expressing the protein of interest, “IPTG”). Spin down (11 krpm, 1 min), remove supernatant, add 100 μL 2XSB + BME, and store at 20°C for further Western blot analysis. Record OD600 of the culture before induction. • Add IPTG to 0.6 mM final concentration (IPTG stock solution 1 M) and grow the cultures for 16 h at 16°C. The inducer concentration and the duration of induction are parameters that can be optimized before large-scale expression. The next day: • Place cultures in ice to chill (approx. 20 min). • While chilling, remove a 1-mL sample from the culture (cells expressing the protein of interest, “+IPTG”). Spin down (11 krpm, 1 min), remove supernatant, add 100 μL 2XSB + BME, and store at 20°C. Record OD600 of the culture after expression induction. • Spin 3 L of culture in a precooled centrifuge with an appropriate rotor for 1-L bottles (3.5 krpm for 15 min). • Pour off supernatant. • Resuspend pellet in 1/50th of original volume with ice-cold lysis buffer A (a 3-L culture cell pellet is resuspended in 60 mL lysis buffer A). The resuspension volume can be adjusted according to the density of cell culture after overnight growth. • Transfer suspended cells in new precooled 50-mL tubes. • Flash freeze resuspended cells in liquid nitrogen. • Store at 80°C. Cells can be stored at 80°C for a few weeks or thawed right away. The flash freezing step is important in order to improve cell lysis. Cell lysis and membrane preparation • Thaw cell pellets in ice water. • Add 600 μL 10 mg/mL lysozyme (100 μg/mL final concentration). • Add 600 μL 100 mM PMSF (1 mM final concentration). • Add 5 μL benzonase. • Incubate on ice for 30 min with periodic vortexing. • Lyse cells using the selected method: high pressure homogenizer Emulsiflex or French press are preferred methods here (prepare extra lysis buffer for cleaning the Emulsiflex chamber). The extract should become clearer. •

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•

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Remove 20 μL and combine with 20 μL 2XSB + BME (cell extract). Transfer cell extract to 50-mL centrifugation tubes. Spin cell extract in a precooled centrifuge with appropriate rotor for 50-mL tubes (3.5 krpm for 15 min). Resuspend pellet in 60 mL ice-cold lysis buffer A. Remove 20 μL and combine with 20 μL 2XSB + BME (unbroken cells and inclusion bodies). Lysis buffer A: 20 mM HEPES pH 7.4, 150 mM NaCl, 5 mM MgCl2, 10% glycerol, 1 mM PMSF. Transfer the supernatant to ice-cold ultracentrifuge tube. Harvest the membranes by spinning the supernatant at 50,000
g for 1 h. After centrifugation: Remove 20 μL of supernatant and combine with 20 μL 2XSB + BME (soluble proteins). Carefully remove the rest of the supernatant from the ultracentrifugation tube. Try to get all the liquid and leave the pellet intact. Add 4.5 mL solubilization buffer B to the pellet and disburse it using a homogenizing pestle. Solubilization buffer B: 50 mM HEPES pH 7.4, 150 mM NaCl, 15% glycerol, 1 mM PMSF. Fully homogenize the disbursed pellet using a homogenizing pestle. Remove 20 μL of suspended pellet and combine with 20 μL 2XSB + BME (membranes). Flash freeze the membranes in liquid nitrogen. The membranes can be stored at 80°C or used right away for solubilization without flash freezing. If suitable solubilization conditions are already known, the whole membrane fraction can be solubilized or aliquots can be prepared (200 μL) in order to perform a detergent screening. All the collected samples can be analyzed by SDS-PAGE followed by a Coomassie blue staining (if the protein is highly abundant) or Western blotting to ensure a proper production and presence in the membrane fraction (Fig. 2A).

2.3 Solubilization Working with membrane proteins in defined in vitro conditions urges to find a detergent, or detergent mix, that retains the protein folded and active

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Fig. 2 E. coli GlpG and human RHBDL2 expression and solubilization conditions screen. (A) Expression of GlpG-His6 and RHBDL2-His6 and membrane preparation from E. coli cells. Expression is induced with 0.6 mM IPTG in the bacterial cell culture, and cells are grown for 16 h at 16°C. Expression and presence of fusion proteins in the membrane fraction are validated by Western blotting using the penta-his antibody (QIAGEN, final concentration 0.2 ng/μL). Black arrowheads correspond to GlpG-His6, and open arrowheads correspond to RHBDL2-His6. (B) Detergent condition screen in order to solubilize GlpG and RHBDL2. Solubilized protein fraction (S) is compared to insoluble fraction (IS) revealing that CHS increases solubility of RHBDL2.

(Kubicek, Block, Maertens, Spriestersbach, & Labahn, 2014; Roy, 2015). Given complex nature of biological membranes, frequently there is no ideal choice. Optimization is entirely empirical and time consuming. This is particularly challenging during purification of membrane proteins, which frequently leads to the loss of essential lipids. Hence, it is often useful to include lipids or lipid derivatives during solubilization. For rhomboid proteases, we previously observed that cholesteryl hemisuccinate (CHS) increases the stability and activity (Lemberg et al., 2005). In this section, we propose a quick and convenient screen to identify a suitable detergent among those most frequently used to solubilize membrane proteases. This solubilization screen can be easily coupled to the activity probing with ActivX TAMRA-FP (Section 3.1) or boron-dipyrromethene (BODIPY) FL casein (Section 3.2).

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Detergent screening • Start from 200 μL aliquots of bacterial membranes. For each of these aliquots, a different detergent or varying detergent concentrations can be tested. • Thaw bacterial membranes in ice water. • For the solubilization of GlpG and RHBDL2, five different detergent conditions are tested (final concentrations are indicated): n-Dodecyl β-D-maltoside (DDM) 1%, Triton X-100 (T-X100) 1%, NP-40 1%, DDM 1%, and CHS 0.05%, Triton X-100 1%, and CHS 0.5%. Stock solutions of detergents should be freshly prepared. CHS (in powder) is dissolved at room temperature for 2 h. Detergent Stock Solutions

Composition

DDM

50 mM HEPES pH 7.4, 150 mM NaCl, 5 mM MgCl2, 10% glycerol, 1 mM PMSF, 2% DDM

T-X100

50 mM HEPES pH 7.4, 150 mM NaCl, 5 mM MgCl2, 10% glycerol, 1 mM PMSF, 2% T-X100

NP-40

50 mM HEPES pH 7.4, 150 mM NaCl, 5 mM MgCl2, 10% glycerol, 1 mM PMSF, 2% NP-40

DDM/CHS

50 mM HEPES pH 7.4, 150 mM NaCl, 5 mM MgCl2, 10% glycerol, 1 mM PMSF, 2% DDM, 0.1% CHS

T-X100/CHS

50 mM HEPES pH 7.4, 150 mM NaCl, 5 mM MgCl2, 10% glycerol, 1 mM PMSF, 2% T-X100 2%, 0.1% CHS

• • • • • •

•

Add 200 μL of the detergent stock solution on top of the membrane aliquot. Solubilize membrane proteins for 1 h on a wheel at 4°C. This step can be extended to 2 h to increase the amount of solubilized proteins. Transfer extract to ice-cold ultracentrifuge tubes. Spin the extract at 90,000
g for 1 h. Remove 20 μL of the supernatant and combine with 20 μL 2XSB + BME (S DDM, S T-X100, etc.) Resuspend the pellet with 400 μL of solubilization buffer (B). Remove 20 μL and combine with 20 μL 2XSB + BME (IS DDM, IS T-X100, etc.). This fraction is the insoluble fraction. All the collected samples from the detergent screen can be analyzed by SDS-PAGE gel electrophoresis followed by a Coomassie blue staining or

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Western blotting to select the best detergent (Fig. 2B). For the large-scale experiment, a DDM/CHS mix is chosen to solubilize GlpG, and a T-X100/CHS mix is selected to solubilize RHBDL2. Large-scale solubilization The method applied to solubilize the membrane fraction of the 3-L culture of E. coli cells is similar to the detergent screen. • Start from 4.5 mL of bacterial membranes (in a 15-mL conical tube). • Thaw bacterial membranes in ice water. • Add 4.5 mL of detergent stock solution on top of the membrane aliquot. GlpG solubilization conditions: 50 mM HEPES pH 7.4, 150 mM NaCl, 5 mM MgCl2, 10% glycerol, 1 mM PMSF, DDM 2%, CHS 0.1%. RHBDL2 solubilization conditions: 50 mM HEPES pH 7.4, 150 mM NaCl, 5 mM MgCl2, 10% glycerol, 1 mM PMSF, T-X100 2%, CHS 0.1%. • Solubilize membrane proteins for 1 h on a rotating wheel at 4°C. • Transfer extract to ice-cold ultracentrifuge tubes. • Spin the extract at 90.000
g for 1 h. • Remove 20 μL of the supernatant and combine with 20 μL 2XSB + BME. • Resuspend the pellet with 9 mL of solubilization buffer B. Remove 20 μL and combine with 20 μL 2XSB + BME. • It is recommended not to flash freeze solubilized proteins at this step but to start the purification process immediately after.

2.4 Purification 2.4.1 Materials and Equipment – Ni-NTA Agarose resin (ABT, USA) – Imidazole (Merck, Germany) – Gravity column (Bio-rad Econo-Pac® Chromatography Column 1.5
12 cm2, Bio-rad, USA) – 2-Way stopcock 2.4.2 Methods • Wash and equilibrate Ni-NTA agarose resin during membrane protein ultracentrifugation (usually, the resin is in a 50% slurry state). • Transfer 2 mL of resin slurry into a 15-mL centrifuge tube.

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Add 10 mL of resin wash buffer: 50 mM HEPES pH 7.4, 500 mM NaCl, 1 mM MgCl2, 10% glycerol, 1 mM PMSF, 5 mM β-Mercaptoethanol. Incubate the resin with wash buffer at 4°C for 5 min. • Spin the resin in a precooled centrifuge (4°C) 1 min at low speed (400 rpm). • Pour off the supernatant without touching the resin and repeat the wash process two more times. • After the last resin centrifugation, pour off as much supernatant as possible. • Transfer the solubilized protein fraction (9 mL) with the resin pellet in the 15-mL conical tube. • Incubate the solubilized proteins with the resin at 4°C on a rotating wheel for 1 h 30 min. The following process has to be done at 4°C: • Transfer the mix in a gravity column. • Let the resin settle down. • Open the stopcock in order to obtain a slow flux (one drop every second) and recover the “flow-through” fraction in a glass beaker. • Use the flow-through to rinse the tube which was used for the previous incubation. • Transfer the flow-through fraction again on the gravity column. To avoid resin resuspension, pour the flow-through onto the side of the column using a glass pipette. • Open the stopcock in order to obtain a slow flux (one drop every second) and recover the “flow-through” fraction in a glass beaker. Remove 20 μL of the flow-through and combine with 20 μL 2XSB + BME. • Pour 20 mL of wash buffer 1 onto the sides of the column: 50 mM HEPES pH 7.4, 500 mM NaCl, 1 mM MgCl2, 30 mM imidazole, 10% glycerol, 0.1% DDM (GlpG), or 0.1% T-X100 (RHBDL2). • Open the stopcock in order to obtain a slow flux (one drop every second) and recover the “Wash 1” fraction in a glass beaker. Remove 20 μL of the wash fraction and combine with 20 μL 2XSB + BME. It is important that the resin never gets dry in the column. The volume of buffer used to wash the resin and imidazole concentration can be adapted to improve the purity of the elution fractions. • Pour 20 mL of wash buffer 2 onto the sides of the column: 50 mM HEPES pH 7.4, 200 mM NaCl, 1 mM MgCl2, 30 mM imidazole, 10% glycerol, 0.1% DDM (GlpG), or 0.1% T-X100 (RHBDL2).

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Open the stopcock in order to obtain a slow flux (one drop every second) and recover the “Wash 2” fraction in a glass beaker. Remove 20 μL of the wash fraction and combine with 20 μL 2XSB + BME. • Add 10 mL of elution buffer onto the column: 50 mM HEPES pH 7.4, 200 mM NaCl, 1 mM MgCl2, 400 mM imidazole, 10% glycerol, 0.1% DDM (GlpG), or 0.1% T-X100 (RHBDL2). • Open the stopcock in order to obtain a slow flux (one drop every second) and recover 1 mL elution fractions into 1.5-mL tubes. Remove 20 μL of every elution fraction and combine with 20 μL 2XSB + BME (elution fraction 1, 2, …). Also remove 20 μL of every fraction to estimate protein concentration. • All the collected samples from the solubilization and purification process can be analyzed by SDS-PAGE followed by Coomassie blue staining to estimate the purity of the elution fractions (Fig. 3). The amount of purified protein can be estimated using BCA or Bradford assay kits. The most abundant fractions can be pooled, aliquoted in small volumes (100 μL), flash frozen in liquid nitrogen, and stored at 80°C. Throughout the production and purification process, it is crucial to retain protease activity, and it therefore required to determine activity at every step of the protocol. The next chapter describes different techniques to probe rhomboid protease activity in the membrane, in a solubilized state and after purification.

3. IN VITRO PROBING OF RHOMBOID PROTEASE ACTIVITY 3.1 Serine Hydrolase Probe to Test Reactivity of Active Site Residues As one of the major bottlenecks is solubilization of the recombinant rhomboid from the membrane fraction, it can be advantageous to optimize expression conditions by validating rhomboid protease activity in the membrane fraction by activity-based protein profiling, which is often used to analyze enzymes in complex proteomic samples (Galmozzi, Dominguez, Cravatt, & Saez, 2014; Kidd, Liu, & Cravatt, 2001). Activity-based profiling makes use of a probe with a reactive group that is able to target and chemically modify the enzyme active site. Serine in the active site of rhomboids efficiently reacts with fluorophosphonate (FP) war heads. Therefore, labeling of the protease with this compound can serve as a direct readout for enzyme activity

Fig. 3 E. coli GlpG and human RHBDL2 purifications by Ni-NTA chromatography. (A) GlpG-His6 is solubilized from membranes using a DDM-CHS detergent mix. The different fractions are run on SDS-PAGE, and the gel is stained with Coomassie blue. From a 3-L culture, the yield is about 1.5 mg of protein. (B) RHBDL2-His6 is solubilized from membranes using a T-X100-CHS detergent mix. The different fractions are loaded on SDS-PAGE, and the gel is stained with Coomassie blue. From a 2-L culture, around 0.5 mg of protein is purified.

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Fig. 4 E. coli GlpG and human RHBDL2 activity monitoring using ActivX TAMRA-FP serine hydrolase probe. (A) The ActivX TAMRA-FP serine hydrolase probe enables selective labeling of active serine hydrolases in both complex samples such as a membrane fraction and in a solubilized state. Fluorophosphonate is coupled to the TAMRA fluorophore. (B) Active site integrity of GlpG and RHBDL2 over solubilization and purifications steps. TAMRA-FP fluorescence is visualized on SDS-PAGE in cell membranes containing GlpG and RHBDL2, in a solubilized state and after purification. Samples are incubated with (100 and 400 μM) or without (0) serine protease inhibitor 3,4-dichloroisocoumarin (3,4-DCI). The labeling specificity is confirmed by Western blotting using the pentahis antibody (QIAGEN, final concentration 0.2 ng/μL).

(Liu, Patricelli, & Cravatt, 1999). FP can be coupled to fluorescent markers such as rhodamine or its derivatives, e.g., tetramethylrhodamine (TAMRA) (Fig. 4A). FP-labeled rhomboids are afterward detected by SDS-PAGE and fluorescent gel scanning. This method has previously been used to label rhomboid proteases either purified or directly in the membrane (Sherratt et al., 2012; Wolf et al., 2015; Xue & Ha, 2012). In this section, we demonstrate the use of the TAMRA-FP molecule to probe GlpG and RHBDL2 activity in different environments during the purification process. 3.1.1 Materials and Equipment – Source material: membrane fraction, solubilized or purified protein fraction

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– ActivX® TAMRA-FP Serine Hydrolase Probe (Catalog number 88318, ThermoFisher, USA) – 3,4-Dichloroisocoumarin (Merck, Germany) – Dimethyl sulfoxide (DMSO) – Water bath/thermomixer – Fluorescent imager (laser excitation/emission wavelengths: 552/575 nm) ImageQuant LAS 4000 (GE Healthcare, United Kingdom) 3.1.2 Methods • Stock solution of TAMRA-FP serine hydrolase probe is prepared according to the manufacturer’s recommendations: e.g., the probe is dissolved in 100 μL of DMSO to prepare a 0.1-mM stock solution. • Membrane fractions, solubilized proteins, and purified GlpG and RHBDL2 are mixed with the probe (final concentration 0.5 μM) in buffer B. 0.5 μg of purified GlpG is sufficient to obtain a strong fluorescence signal. Because of their complex composition a higher amount of membrane fraction and solubilized protein has to be used to obtain a comparable fluorescence signal. The total volume of each reaction is 20 μL. A serine protease inhibitor, 3,4-dichloroisocoumarin, is added to the mix prior to the probe in variable amounts (final concentration 0, 100, or 400 μM, respectively). • Samples are incubated at 37°C in a thermomixer (without agitation) for 1 h and protected from light. • The reaction is stopped by addition of 20 μL 2XSB + BME. • Samples are subjected to SDS-PAGE, and fluorescence signal is visualized using a fluorescent imager with Cy3 filters and green laser (excitation wavelength 552 nm and emission wavelength 575 nm) (Fig. 4B). Proteins are then blotted on PVDF membranes and visualized with antiHis antibodies.

3.2 EnzChek Protease Assay for Efficient Screening of Rhomboid Protease Activity EnzChek protease assay is a fluorescence-based method for detecting and measuring protease activity. The substrate is a casein derivative, labeled with a green-fluorescent BODIPY FL dye. Highly fluorescent BODIPY FL peptides are released after casein hydrolysis by a protease (Fig. 5A). The emitted fluorescence can be measured using a microplate reader and correlates to the protease activity. The assay is featured by a high sensitivity and pH insensitivity.

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Fig. 5 BODIPY FL casein cleavage by E. coli GlpG. (A) In unprocessed state, BODIPY FL casein is quenched and only emit a basal fluorescence. Once incubated with a protease, the casein is cleaved and leads to fluorescent cleavage products. (B) BODIPY FL casein is mixed with GlpG (5 μg) without or with 3,4-dichloroisocoumarin (3,4-DCI; concentration as indicated). Fluorescence is measured over time. Prolonged incubation with GlpG leads to an increase of fluorescence.

The simple and fast setup of the assay allows it to be applied in highthroughput screenings. Despite relying on a soluble substrate, it has been successfully used to study E. coli GlpG protease activity (Wang et al., 2006) and for kinetic studies (Lazareno-Saez et al., 2013). 3.2.1 Materials and Equipment – Purified protein fraction – EnzChek® Protease assay kit (Catalog number E6638, Thermo Fisher Scientific, USA) – 96-Well microplate, flat and transparent bottom – Fluorescence microplate reader (Laser Ex/Em: 485/530 nm) Tecan Infinite M1000 Pro (Tecan, Switzerland) 3.2.2 Methods • 1
Digestion buffer and 5 μg/mL working solution of BODIPY casein are prepared according to the manufacturer’s protocol.

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Dilute 0.5 μg of purified protein (e.g., GlpG) in buffer containing 50 mM HEPES pH 7.4, 200 mM NaCl, 10% glycerol, 0.1% DDM to a final volume of 100 μL. 3,4-Dichloroisocoumarin is added to a final concentration of 100 or 400 μM. Add 100 μL of working solution of BODIPY casein (final concentration 2.5 μg/mL). The total volume of each reaction mix is 100 μL. It is supplemented by 100 μL of BODIPY casein working solution before starting fluorescence measurement. Reactions are performed in 96-well microplate. A negative control mix composed of 100 μL buffer and 100 μL BODIPY casein working solution (buffer-only control) is included. Set up the fluorescence microplate reader: excitation wavelength ¼485  12.5 nm, emission wavelength ¼ 530  15 nm. Start the fluorescence measurement over time (37°C for 4 h) (Fig. 5B).

3.3 Rapid Testing of In Vitro-Generated TM Domain Substrates One aim of the in vitro study of intramembrane protease is to be able to directly monitor the cleavage of known or candidate substrates. In vitro experiments allow recording reaction kinetics and studying required intrinsic features for a membrane protein to be cleaved. Setting up a robust in vitro cleavage assay requires reconstituting a minimal system and therefore to have, on one hand, rhomboid in an active state (in the membrane, in a solubilized form or purified) and on the other hand substrates. As said previously, it can be highly challenging and time consuming to purify TM protein substrates because expression, solubilization, and purification protocols have to be adjusted for each protein. Thus, we have set up an efficient method to generate radiolabeled TM substrate peptides allowing quick testing of cleavage in a crude cytoplasmic fraction of the wheat germ-derived in vitro translation system. This method was successfully applied to a broad range of rhomboid proteases and substrates (Lemberg et al., 2005; Strisovsky et al., 2009). In brief, the coding region of interest is amplified by a standard PCR reaction thereby offering a very flexible substrate design (Lemberg & Martoglio, 2003). As forward primer, an oligonucleotide containing the SP6 or T7 promoter, Kozak initiation sequence, ATG for initiation, followed by the respective coding region is used. As reverse primer an oligonucleotide 50 -N6CTAN20-30 of which the N20 region is complementary to the cDNA introduces a TAG stop codon at the desired position of the open reading frame. Subsequently, capped

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mRNA is generated by in vitro transcription using SP6 or T7 RNA polymerase. Radiolabeled TM peptides are generated by cell-free translation using commercial wheat germ extract. Prior the cleavage assay, translation is either stopped by addition of EGTA (50 mM) or by applying puromycin (1 μM). 3.3.1 Materials and Equipment – DNA template to PCR amplify the substrate’s TMD (cDNA, plasmid) – Forward and reverse primers: the forward primer includes the SP6 promoter, Kozak initiation sequence, ATG for initiation followed by coding region of interest. The reverse primer has to contain a TAG stop codon. – Pfu DNA polymerase or any other polymerase with proof reading activity – Phenol – Phenol/chloroform – Chloroform – Sodium acetate (NaOAc) – SP6 RNA polymerase – m7G(50 )ppp(50 )G RNA CAP structure analog (New England Biolabs, USA) – ATP, CTP, GTP, UTP – Spermidine – RNasin® Ribonuclease Inhibitor (Promega, USA) – Magnesium acetate (Mg(OAc)2) – Dithiothreitol (DTT) – Wheat germ extract (Promega, USA) – Potassium acetate (KOAc) – Amino acid mixture minus methionine – [35S]-methionine – Ethylenediaminetetraacetic acid (EDTA) – Puromycin – EGTA (ethylene glycol-bis (β-aminoethyl ether)-N,N,N0 ,N0 -tetraacetic acid) – Vacuum gel dryer – Tris-Bicine gel system and buffers – Typhoon FLA 7000 (GE Healthcare, United Kingdom) 3.3.2 Methods • PCR: the reaction is performed in a total volume of 100–200 μL according to the manufacturer’s description.

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Phenol/chloroform extraction: add one volume of phenol, vortex 10 s, and centrifuge 1 min at full speed. Transfer the aqueous phase in a new tube. Add one volume of phenol/chloroform mix, vortex 10 s, and centrifuge 1 min at full speed. Transfer the aqueous phase in a new tube. Add one volume of chloroform, vortex 10 s, and centrifuge 1 min at full speed. Transfer the aqueous phase in a new tube. Ethanol precipitation of DNA: add a 1/10th volume of a 3-M NaOAc (pH 5.2) solution and 2.5 volumes of ice-cold absolute ethanol. Vortex shortly, spin 5 min at full speed, and discard supernatant. Wash the DNA pellet with approximately 180 μL of ice-cold 70% ethanol, spin 5 min at full speed, and discard supernatant. Dry the pellet by gentle warming (with the tube lid open) and dissolve in 50 μL H2O. Transcription to obtain 50 -capped RNA: the reaction is performed in a total volume of 100 μL. DNA (50 μL of the DNA previously prepared), 100 U SP6 RNA polymerase, 40 mM Tris–HCl pH 8.0, 500 μM m7G (50 )ppp(50 )G RNA CAP structure analog, 2 mM ATP, 2 mM CTP, 2 mM GTP, 2 mM UTP, 1 mM spermidine (optional), 40 U RNasin (optional), 20 mM Mg(OAc)2, 5 mM DTT. The reaction is performed at 42°C for 2 h. Phenol/chloroform extraction: same protocol as described earlier. Ethanol precipitation of transcript: add a 1/10th volume of a 3-M NaOAc (pH 5.2) solution, add 2.5 volumes of ice-cold absolute ethanol. Vortex shortly, spin 5 min at full speed, and discard supernatant. Wash the RNA pellet with approximately 180 μL of ice-cold 70% ethanol, spin 5 min at full speed, and discard supernatant. Dry the pellet by gentle warming (with the tube lid open) and dissolve in 20 μL H2O. To analyze the quality of the RNA, 0.5 μL of RNA mixed with 6 μL of RNA-buffer (1
) can be heated at 56°C for 3 min and loaded on an agarose gel. Translation: the reaction is performed in a total volume of 25 μL. 0.5 μL RNA, 10 μL wheat germ extract, amino acid mixture minus methionine (20 μM each amino acid), 1 μL [35S]-methionine. The translation reaction can be improved changing magnesium and potassium concentrations and the amount of RNA. Translation is either stopped by addition of EGTA (50 mM) or by applying puromycin (1 μM). Samples can be directly used in an in vitro cleavage assay or flash frozen and stored at 80°C. Cleavage assay: the reaction is performed in a total volume of 40 μL. 2 μL of the translation mixture is added to 38 μL of buffer containing the

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rhomboid protease (e.g., 5 μg of purified protein) in 50 mM HEPES/ NaOH (pH 7.4), 10% glycerol, and 50 mM EDTA. The reaction is incubated at 30°C for 1 h (the duration of incubation and the amount of rhomboid can be adjusted). Protein precipitation: proteins are next precipitated with 10% trichloroacetic acid and then centrifuged for 2 min (10 krpm). Supernatant is discarded. Samples are washed with 100 μL acetone, and centrifugation is repeated. After removing the acetone, the pellet is dissolved in SDSsample buffer (30 μL) by incubation at 65°C for 15 min. Samples are then loaded on Tris-Bicine polyacrylamide gels (Wiltfang, Arold, & Neuhoff, 1991) or any other suitable gel system such as BisTris/MES gels. Following electrophoresis, gels are incubated in 40% methanol, 10% acetic acid for 30 min to allow fixation of polypeptides. Gels are then placed on a wet Whatman paper and dried under vacuum at 65°C for 1 h. Labeled proteins are visualized by filmless autoradiography and scanned using a Typhoon FLA 7000.

3.4 Cleavage of Chimeric or Tagged Substrates To study rhomboid-catalyzed cleavage in a defined setup, cleavage assays consist of purifying a tagged version of the potential substrates. Instead of expressing the entire protein, which would lead to a long process of optimization as discussed before, a chimeric genetic construct can be used. Therefore, a gene fragment encoding the substrate TM domain is cloned into the expression vector pET25b(+) and by this tagged with the bla gene (encoding the β-lactamase protein) and bla signal sequence and a sequence encoding a His6 tag (Fig. 6A). The signal sequence of the resulting fusion protein is supposed to be cleaved by a signal peptidase once inserted in the membrane: the TM domain corresponding to the substrate is supposed to be cleaved by the rhomboid protease. The ability of rhomboids to cleave such chimeric substrates was validated in several studies, where a TM domain of the multispanning protein LacY was fused to the secreted proteins β-lactamase and MBP (maltosebinding protein) (Akiyama & Maegawa, 2007; Maegawa, Ito, & Akiyama, 2005; Maegawa, Koide, Ito, & Akiyama, 2007). The cleavage of tagged substrates was also used to monitor the activity of other intramembrane proteases such as γ-secretase complex (Fukumori & Steiner, 2016). In this study, amyloid precursor protein-tagged proteins were solubilized from E. coli membranes using a urea containing

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Fig. 6 Cleavage of chimeric substrates. (A) Gurken chimeric construct. The fusion protein is composed of a signal sequence, beta-lactamase, Gurken TM domain, and a His6 tag. (B) Gurken chimeric substrate (CS) urea solubilization and purification by Ni-NTA chromatography. Gray arrowhead corresponds to Gurken CS. (C) Gurken CS cleavage by GlpG. The cleavage is monitored by Western blotting using the penta-his antibody (QIAGEN final concentration 0.2 ng/μL). Gray arrowhead corresponds to the full-length version of Gurken CS, and the star (*) points out the cleavage fragment containing the His6 tag. Black arrowhead indicates GlpG-His6. GlpG is incubated with Gurken CS and an increasing concentration of 3,4-dichloroisocoumarin (0, 100, and 400 μM), a serine protease inhibitor, to ensure cleavage specificity.

lysis buffer. The same method is used here to solubilize chimeric rhomboid substrates. To illustrate this approach, a chimeric protein containing the TM domain of Gurken, a Drosophila growth factor cleaved by rhomboid proteases (Urban, Lee, & Freeman, 2002), was expressed, solubilized, and purified from E. coli cells.

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3.4.1 Materials and Equipment – Urea 3.4.2 Methods Expression, solubilization, and purification • The expression protocol is similar to the one described in Section 2.2 but the solubilization method differs. • Starting from membrane pellet after ultracentrifugation, the resuspension is directly achieved in 10 mL urea solubilization buffer: 20 mM Tris pH 8.5, 6 M urea, 1% Triton X-100, 1% SDS, 1 mM CaCl2, 150 mM NaCl. • Incubate membranes in solubilization buffer overnight on a wheel at 4°C. • The next day, ultracentrifuge the solubilization mix (50,000
g, 15 min, 4°C). • Remove 20 μL of the supernatant and combine with 20 μL 2XSB + BME. This corresponds to urea solubilized membrane proteins (solubilized fraction). • Resuspend the insoluble pellet with 10 mL of solubilization buffer B (refer to Section 2.2). Remove 20 μL of the supernatant and combine with 20 μL 2XSB + BME. This corresponds to the “insoluble fraction.” • Transfer the solubilized protein fraction (10 mL) onto the Ni-NTA resin pellet (corresponding to a 2-mL slurry sample) in a 15-mL conical tube (refer also to Section 2.2 for resin wash and preparation). • Incubate the solubilized proteins/resin mix on a wheel at 4°C for 1 h 30 min. • The purification protocol is similar to the one described in Section 2.2, but the washing buffers are different: Washing buffer 1: 20 mM Tris pH 8.5, 1% Triton X-100, 300 mM NaCl. Washing buffer 2: 20 mM Tris pH 8.5, 0.2% SDS, 300 mM NaCl. Elution buffer: 20 mM Tris pH 8.5, 0.2% SDS, 300 mM NaCl, 100 mM imidazole. • All collected samples from the solubilization and purification process can be analyzed by SDS-PAGE gel electrophoresis followed by a Coomassie blue staining (Fig. 6B). The purity of elution fractions could be improved by adding an increasing concentration of imidazole during the washing steps.

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Cleavage assay • The total volume of each cleavage mix is 20 μL. Approximately 0.5 μg of GlpG is incubated with 2 μg of Gurken elution fraction at 37°C for 1 h in buffer B supplemented with 0.1% DDM. If necessary, 3,4dichloroisocoumarin is added to the mix (final concentration 100 or 400 μM, respectively). • Stop the reaction by addition of 20 μL 2XSB + BME. • Mixtures are then loaded on a 15% acrylamide SDS-Gel and analyzed by Western blotting (Fig. 6C).

ACKNOWLEDGMENTS We thank Josephine Bock for technical assistance and Verena Dederer, Nathalie K€ uhnle, and Julia Knopf for critical reading of the manuscript. The work was supported by the Deutsche Forschungsgemeinschaft (FOR2290, TP1).

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CHAPTER FIVE

Expression, Purification, and Enzymatic Characterization of Intramembrane Proteases R. Zhou1, Y. Shi1, G. Yang1 Ministry of Education Key Laboratory of Protein Science, Tsinghua-Peking Joint Center for Life Sciences, Beijing Advanced Innovation Center for Structural Biology, School of Life Sciences, Tsinghua University, Beijing, China 1 Corresponding authors: e-mail address: [email protected]; [email protected]; [email protected]

Contents 1. Introduction to Intramembrane Proteases 2. Expression and Purification of GlpG, S2P, and PSH 2.1 Glimpse of Rhomboid, S2P, and PSH 2.2 Expression of GlpG and S2P 2.3 Sequence-Based Protein Engineering and Expression of PSH 2.4 Purification of GlpG, S2P, and PSH 3. Enzymatic Activity Assays for Intramembrane Proteases 4. Expression and Purification of Human γ-Secretase 4.1 Introduction to Human γ-Secretase 4.2 Design of the Mammalian Expression Vector pMlink 4.3 Transient Expression of Human γ-Secretase 4.4 Purification of Human γ-Secretase 5. Enzymatic Activity Assay for Human γ-Secretase 6. Structures of I-CLiPs 7. Concluding Remarks Acknowledgments References

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Abstract Intramembrane proteases catalyze peptide bond hydrolysis in the lipid bilayer and play a key role in numerous cellular processes. These integral membrane enzymes consist of four classes: site-2 protease (S2P), rhomboid serine protease, Rce1-type glutamyl protease, and aspartyl protease exemplified by presenilin and signal peptide peptidase (SPP). Structural elucidation of these enzymes is important for mechanistic understanding of their functions, particularly their roles in cell signaling and debilitating diseases such as Parkinson’s disease and Alzheimer’s disease. In the past decade, rigorous effort has led to determination of the crystal structures of S2P from archaebacterium, rhomboid serine

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protease from E. coli (GlpG), and presenilin/SPP from archaebacterium (PSH). A novel method has been developed to express well-behaved human γ-secretase, which facilitated its structure determination by cryoelectron microscopy (cryo-EM). In this chapter, we will discuss the expression and purification of intramembrane proteases including human γ-secretase and describe the enzymatic activity assays for these intramembrane proteases.

1. INTRODUCTION TO INTRAMEMBRANE PROTEASES Regulated intramembrane proteases (RIP or intramembrane-cleaving proteases, I-CLiPs) execute proteolysis in lipid bilayer environment (Brown, Ye, Rawson, & Goldstein, 2000). On the basis of catalytic mechanisms, the I-CLiPs can be divided into four major types: rhomboid serine protease, Rce1-type glutamyl protease, zinc-bound site-2 protease (S2P), and aspartyl protease exemplified by presenilin and signal peptide peptidase (SPP) (Ebinu & Yankner, 2002; Erez, Fass, & Bibi, 2009; Langosch, Scharnagl, Steiner, & Lemberg, 2015; Urban & Freeman, 2002; Wolfe & Kopan, 2004). Regulated intramembrane proteolysis was first observed in the study of sterol homeostasis. S2P plays a key role in the sterol regulatory elementbinding protein (SREBP) pathway that regulates sterol level in cells (Brown & Goldstein, 1997; Rawson et al., 1997). Low cholesterol level prompts SREBP translocation from ER to Golgi, where SREBP is sequentially cleaved by the site-1 protease (S1P) and S2P to release the transcriptional activation domain at the N-terminus. S2P is also found to cleave ATF6, a membrane-bound transcription factor involved in ER stress response (Ye et al., 2000). The rhomboid protease was first identified in the study of epidermal growth factor receptor signaling in Drosophila and soon found to play a critical role in pathophysiological events such as Parkinson’s disease (PD) (Bang & Kintner, 2000; Meissner, Lorenz, Weihofen, Selkoe, & Lemberg, 2011; Wasserman, Urban, & Freeman, 2000). Presenilin was identified through analysis of early onset familial AD (FAD) patients (Sherrington et al., 1995). Presenilin is usually incorporated into γ-secretase to catalyze cleavage of type I transmembrane proteins such as amyloid precursor protein (APP) (Li, Lai, et al., 2000; Li, Xu, et al., 2000). Cleavage of APP results in the generation of amyloid-β peptides, which through aggregation form amyloid-β plaques in the human brain (Wolfe et al., 1999). Presenilins also have other substrates such as Notch and

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N-cadherin (De Strooper et al., 1999). Over 200 AD derived mutations have been mapped to presenilin-1. In 2002, SPP was discovered as a presenilin-type aspartyl protease which has the opposite topological arrangement compared to presenilin (Weihofen, Binns, Lemberg, Ashman, & Martoglio, 2002). The Rce1-type glutamyl protease was recently identified as a new class of intramembrane protease (Manolaridis et al., 2013; Pryor et al., 2013). For the I-CLiPs (except Rce1-type glutamyl proteases), substrate cleavage occurs in the lipid membrane, which is a hydrophobic environment. However, water is required for peptide bond hydrolysis. This quagmire raises the key question of how water molecules gain access to the proteolysis active site. Another question is how the substrate is accessible to the active site. In addition, the involvement of I-CLiPs in disease development makes them ideal targets for potential therapeutic intervention. Taken together, detailed structural information of I-CLiPs is a prerequisite for mechanistic understanding of their functions. In the past decade, structural information on S2P, rhomboid, and PSH has become available. More recently, we have successfully expressed human γ-secretase and determined its atomic structure by cryo-EM. These structures have provided significant insights into the cleavage mechanism of I-CLiPs and serve as a rational basis for drug development.

2. EXPRESSION AND PURIFICATION OF GlpG, S2P, AND PSH 2.1 Glimpse of Rhomboid, S2P, and PSH Genetic screening led to identification of a number of I-CLiPs called rhomboid-like proteins (Freeman, 2014; Lemberg & Freeman, 2007). The well-studied rhomboid homolog Rhomboid-1 in Drosophila cleaves the EGF-like ligand Spitz to activate the EGFR signaling (Lee, Urban, Garvey, & Freeman, 2001; Urban, Lee, & Freeman, 2002). Function of the rhomboid family is conserved from Drosophila to mammals in the cleavage-triggered activation of their substrates EGFR, EGF, and ephrinB3 (Freeman, 2014). RHBDL4, a rhomboid in mammals, is thought to play a role in the ER-associated degradation of unfolded protein response (Fleig et al., 2012). A human mitochondrial protein named “presenilin-associated rhomboid like” contributes to PD and type-2 diabetes (Meissner et al., 2011). A complex negative feedback process called the SREBP pathway regulates sterol homeostasis in cells. SREBP is a hairpin-like protein in the ER membrane, with two transmembrane helices and two cytosolic domains

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(Hua, Sakai, Ho, Goldstein, & Brown, 1995; Nohturfft, Brown, & Goldstein, 1998). When cells are deprived of sterol, SREBP together with SREBP cleavage-activating protein are translocated to the Golgi, where SREBP undergoes two sequential cleavages by S1P and S2P to release its N-terminal transcription activation domain for transcriptional stimulation of the genes that are responsible for de novo synthesis and uptake of fatty acids and sterol (DeBose-Boyd et al., 1999; Duncan, Dave, Sakai, Goldstein, & Brown, 1998; Nohturfft, DeBose-Boyd, Scheek, Goldstein, & Brown, 1999; Sakai, Nohturfft, Goldstein, & Brown, 1998; Sakai, Rawson, et al., 1998). Apart from SREBP, another integral transmembrane protein ATF6, which is involved in the unfolded protein response, is also cleaved by S2P (Ye et al., 2000). S2P usually contains one or two PDZ domains that may function in the recognition of substrates (Harris & Lim, 2001). Another feature of S2P is a consensus HExxH sequence at the N-terminus, among which the two histidine residues coordinate a zinc atom, and the glutamate residue activates a water molecule for peptide bond hydrolysis (Rawlings & Barrett, 1995). Much attention is drawn to the aspartyl intramembrane protease, particularly presenilin due to its close relationship with the early onset of FAD. At the time of manuscript preparation, 243 patient-derived mutations have been identified in presenilin-1, and 48 missense mutations are mapped to presenilin-2 (http://www.Alzforum.org). Another important aspartyl protease is SPP, which was identified to cleave signal peptide (McLauchlan, Lemberg, Hope, & Martoglio, 2002; Weihofen et al., 2002). SPP-like family proteases (SPPL), including SPPL2a/b/c, SPPL3, and SPPL4, play distinct roles in cells (Grigorenko, Moliaka, Soto, Mello, & Rogaev, 2004; Ponting et al., 2002; Weihofen et al., 2002). Specifically, SPPL2a/b cleaves tumor necrosis factor alpha (TNF-α), Bri2, and Fas, whereas SPPL3 is mainly involved in the protein glycosylation and Golgi network (Fluhrer et al., 2006; Friedmann et al., 2006; Kirkin et al., 2007; Kuhn et al., 2015; Martin et al., 2008; Voss et al., 2014).

2.2 Expression of GlpG and S2P Rhomboid was the first I-CLiP successfully targeted for structural investigation by a number of independent laboratories. In our case (Wu et al., 2006), 60 prokaryotic rhomboid homologs were selected for ORF subcloning into pET15b (Novagen) and small-scale expression test in E. coli C43 (DE3). After preliminary screening, the E. coli homolog GlpG emerged

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as a clear favorite due to its high yield, good solution behavior, and robust enzymatic activity. The E. coli cells transformed by a GlpG-expressing plasmid were induced with 0.5 mM isopropyl-β-D-thiogalactopyranoside (IPTG) at OD600 of 1.5 and harvested after 8 h (Wu et al., 2006). During purification of the fulllength GlpG, some degradation bands below 10 kDa persisted, suggesting the presence of flexible sequences within the protein. Limited proteolysis was employed to identify a stable core domain amenable for crystallization with different proteases, including trypsin, chymotrypsin, elastase, and subtilisin. Chymotrypsin digestion of GlpG yielded a doublet that both began at residue 87; hence, a core domain (residues 87–276) was engineered, which retained the full proteolytic activity compared to the wild-type protein. Other groups have reported the crystal structure of GlpG using nearly the same expression and purification strategy (Ben-Shem, Fass, & Bibi, 2007; Lemieux, Fischer, Cherney, Bateman, & James, 2007; Wang, Zhang, & Ha, 2006). For S2P, a similar working scheme was adopted. Briefly, 40 S2P homologs were chosen from bacterial and archaeal species, and the ORFs were subcloned into pET15b and pET21b (Novagen), and overexpressed in E. coli BL21 (DE3) and C43 (DE3). After extensive screening, a S2P homolog from Methanocaldococcus jannaschii (mjS2P) was found to exhibit high expression levels and relatively good solution behavior. At OD600 of 1.0–1.2, the E. coli cells were induced by 0.2 mM IPTG at 23°C for 16 h. Some degradation bands showed up during purification. A 25-kDa stable core domain (residues 1–244) was identified by multiple methods including limited proteolysis, mass spectrometry, N-terminal sequencing, and secondary structural prediction (Feng et al., 2007).

2.3 Sequence-Based Protein Engineering and Expression of PSH Because screening of numerous eukaryotic presenilin-1 homologs only led to limited protein production, we sought its archaeal counterparts instead. PSH was selected from 13 archaeal candidates, subcloned into pET21b (Novagen), and overexpressed in BL21 (DE3). At OD600 of 1.5, 0.2 mM IPTG was added to induce protein overexpression at 22°C. Cells were harvested after 16 h and homogenized in lysis buffer containing 25 mM Tris–Cl, pH 8.0, and 150 mM NaCl for further purification. Although a reasonable yield of PSH can be achieved, the wild-type protein is extremely prone to aggregation and precipitation.

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Protein engineering for an optimal PSH variant was designed as follows (Fig. 1A). In the very beginning, sequences of 13 archaeal homologs including PSH were aligned. A number of residues are highly conserved in other archaeal homologs except in PSH. We reasoned that replacement of these residues by the conserved ones might improve the solution behavior of PSH. Twenty PSH variants, each containing one or two missense mutations, were constructed and individually examined for their enzymatic activities. Five mutations, each associated with improved protein solution behavior and robust protease activity, were combined to generate 10 PSH variants, each with two or three mutations. Of these variants, four were further combined to generate six constructs, each with four or five mutations. In the end, the variant bearing five missense mutations, D40N, E42S, A147E, V148P, A229V, was used for crystallization, and most of these mutations affect PSH

Fig. 1 Sequence-based protein engineering and purification of PSH. (A) A work-flow diagram of sequence-based protein engineering for PSH. This approach was designed to improve the solution behavior of PSH. Sequences from 13 archaeal homologs of presenilin-1/SPP are aligned. The residues that are highly conserved in other species but not in PSH are replaced by the conserved ones. The mutations with better behavior and robust activity are combined for further selection and optimization. (B) Purification of the engineered PSH with five mutations and a loop deletion. A representative chromatogram from gel filtration is shown in the upper panel. The peak fractions of gel filtration are visualized on SDS-PAGE in the lower panel. The protein is cleaved into NTD and CTD by V8 protease.

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surface loops. Limited proteolysis was employed to eliminate the surface flexible regions, and V8 protease removed residues 182–209, resulting in the separation of the N-terminal domain (NTD) and C-terminal domain (CTD) (Fig. 1B). The best crystals were derived from the V8-digested protein with five missense mutations described earlier. The strategy of sequence-based engineering may be generally applied to those proteins with bad solution behavior (Li, Dang et al., 2013).

2.4 Purification of GlpG, S2P, and PSH 2.4.1 Required Materials The following protocol will require these materials. 1. Reagents Lysis buffer (25 mM Tris–HCl, pH 8.0, 150 mM NaCl) Imidazole, pH 8.0 Phenylmethylsulfonyl fluoride, Amresco (PMSF, 0754; Solon, OH) 100 mM PMSF solution (in 100% ethanol) n-Decyl-β-D-Maltopyranoside, Anatrace (DM, D322; Maumee, OH) n-Dodecyl-β-D-Maltopyranoside, Anatrace (DDM, D310; Maumee, OH) n-Nonyl-β-D-Maltopyranoside, Anatrace (NM, N330; Maumee, OH) n-Nonyl-β-D-Glucopyranoside, Anatrace (NG, N324; Maumee, OH) n-dodecyl-N, N-Dimethyldodecylamine-N-oxide, Anatrace (LDAO, D360; Maumee, OH) Thrombin, Sigma (T4648; St. Louis, MO) 2. Equipment G-560E Vortex, Scientific Industries (Bohemia, NY) T10 basic ULTRA-TURRAX disperser, IKA EmulsiFlex-C3 homogenizer, Avestin (French press) Avanti™ centrifuge J-25, Beckman Coulter (Palo Alto, CA) Optima™ L-90K Ultracentrifuge, Beckman Coulter (Palo Alto, CA) Type 70 Ti rotator, Beckman Coulter (Palo Alto, CA) 50-mL PP centrifuge tubes, conical bottom with plug seal cap, Corning (China) QB-208 Multipurpose shaker, Kylin-Bell (Haimen, Jiangsu, China) Size-exclusion chromatography system, GE Healthcare (Uppsala, Sweden) Ni2+-NTA affinity column, Qiagen (30210; Hilden, Germany) Superdex-200, GE Healthcare (17517501; Uppsala, Sweden) Econo-pac chromatography columns, Bio-Rad (Hercules, CA)

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2.4.2 Purification of GlpG, S2P, and PSH The general protocol for purification of these proteases is as follows (Fig. 2A and B): Step 1: Homogenize the E. coli cells in lysis buffer as mentioned earlier, supplemented with PMSF to a final concentration of approximately 1 mM. Step 2: Disrupt the cells by French press with two passes at a maximum pressure of 15000 psi. Step 3: Apply the disrupted cells to centrifugation at 23,000
g for 10 min to remove cell debris. Step 4: Collect the supernatant and apply to ultracentrifugation at 150,000
g for 1 h to pellet cell membranes. Step 5: Incubate the membrane fraction with specific detergents for 1 h at 4°C after resuspending the membrane by disperser. Specifically, extract GlpG by 2% (w/v) NG, PSH by 2% (w/v) NM, and S2P by 1% (w/v) DM. Step 6: Subject the extraction to ultracentrifugation at 150,000
g for 30 min to remove undissolved fraction. Harvest the supernatant that contains the solubilized protease for affinity purification. Step 7: Load the supernatant onto Ni2+-NTA affinity column and wash with 25 mM Tris–HCl, pH 8.0, 150 mM NaCl, 20 mM imidazole, and

Fig. 2 Expression and purification of the I-CLiPs. (A) The expression and purification scheme for prokaryotic homologs of I-CLiPs. (B) A generalized flowchart diagram of the purification procedure.

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detergents. Specifically, 0.4% (w/v) NG is used for GlpG, 0.2% (w/v) DM for S2P, and 0.6% (w/v) NM for PSH. Step 8: Elute GlpG from the Ni2+-NTA column with 25 mM Tris–HCl, pH 8.0, 150 mM NaCl, and 250 mM imidazole with 0.4% (w/v) NG. The elution buffer for PSH is identical to that for GlpG except the detergent (0.6% (w/v) NM). Different from GlpG and PSH, S2P is released from the affinity column by thrombin digestion in 25 mM Tris–HCl, pH 8.0, 150 mM NaCl, and 0.2% (w/v) DM. Step 9: Concentrate the eluted protein to 15–20 mg mL–1 and apply to gel filtration (Superdex-200, GE Healthcare) in its elution buffer. The gel filtration buffer for GlpG contains 10 mM Tris–HCl, pH 8.0, 150 mM NaCl, and 0.4% (w/v) NG. S2P is eluted by 10 mM Tris– HCl, pH 8.0, 150 mM NaCl, and 0.15% (w/v) DM. PSH is in the buffer of 25 mM Tris–HCl, pH 8.0, 150 mM NaCl, 0.2% (w/v) NG, and 0.023% (w/v) LDAO.

3. ENZYMATIC ACTIVITY ASSAYS FOR INTRAMEMBRANE PROTEASES Structural biology bridges protein structure with its function. In the case of I-CLiPs, enzymatic activity assays serve as a powerful tool for the examination of structure–function relationship. The enzymatic activity assay of GlpG was reconstituted with an artificial substrate CED4, which is a membrane-associated protein (Chen et al., 2000), in the buffer containing 25 mM Tris–HCl, pH 8.0, 150 mM NaCl, and 0.4% (w/v) NG at 37°C. The final concentrations were 0.1 mg mL–1 for GlpG and approximately 2.0 mg mL–1 for the substrate protein. The reaction was stopped by addition of SDS loading buffer, and all the cleavage products were analyzed by SDS-PAGE. The proteolytic activity of the transmembrane core domain of GlpG used for crystallization was comparable to that of the full-length protein as measured by the extent of CED4 cleavage. Another enzymatic activity assay was performed on the physiological substrate C100Spitz-Flag or a control substrate C100-Flag (Urban & Wolfe, 2005). Both GlpG and its transmembrane core domain cleaved C100Spitz-Flag, but not C100-Flag, indicating the specific nature of cleavage (Wu et al., 2006). On the basis of distinct GlpG structures captured in the fully open, partially open, and closed states (Ben-Shem et al., 2007; Lemieux et al., 2007; Wang & Ha, 2007; Wu et al., 2006), lateral gating by TM5 was purposed to

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regulate substrate entry (Wu et al., 2006) (Fig. 3A). Cross-linking between pairs of residues on TM2 and TM5 (Fig. 3B) resulted in a marked reduction of proteolytic activity; increasing TM5 flexibility led to enhanced proteolytic cleavage (Baker, Young, Feng, Shi, & Urban, 2007). Through kinetic characterization of substrate cleavage on liposomes, substrate gating was shown to be a rate-limiting step (Dickey, Baker, Cho, & Urban, 2013). The membrane protein CED9, an antiapoptosis member of the Bcl-2 family (Hengartner & Horvitz, 1994), was used as artificial substrate in the S2P cleavage assay. The reaction was performed in a buffer containing 10 mM Tris–Cl, pH 8.0, 100 mM NaCl, and 0.2% (w/v) DM at room temperature for 30 min. The concentrations of the enzyme and substrate in the assay were 0.1 and 1.0 mg mL–1, respectively. These assays demonstrate that mutations of the active site residues compromised the proteolytic activity of S2P (Feng et al., 2007). The proteolysis assay for PSH was performed at 37°C for 8 h in the PBS buffer containing 0.02% (w/v) DDM and 20 mM citrate, pH 5.3 (Li, Dang,

Fig. 3 The lateral gating mechanism of GlpG and a representative enzymatic assay for I-CLiPs. (A) The lateral gating mechanism of GlpG. Cross-linking of specific residues on TM2 and TM5 may impede substrate entry and thus result in decreased protease activity. (B) The experimental design to test the lateral gating mechanism. Pairs of specific residues for cross-linking on TM2 and TM5 are shown in the same color. The red region indicates the active site of GlpG. (C) The proteolytic activity of I-CLiPs is examined by substrate cleavage. Briefly, the enzyme and substrate are incubated in a specific reaction buffer. The cleavage products are visualized on an SDS-PAGE gel by Coomassie blue staining. Shown here is an example of substrate cleavage by PSH.

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et al., 2013), using Gurken as the substrate (Fig. 3C) (Torres-Arancivia et al., 2010). Because PSH is an archaeal homolog of presenilin-1, the catalytic subunit of γ-secretase, the proteolysis assay was probed by a specific γ-secretase inhibitor N-[[[(2R,3S)-3-[[(1,1-dimethylethoxy)carbonyl]amino]-2hydroxy-4-phenylbutyl](phenylmethyl) amino]carbonyl]-L-leucyl-L-valine methyl ester WPE-III-31-C (hereafter as III-31-C, Sigma-Aldrich; C0619; St. Louis, MO). The γ-secretase modulator GSM-1 was also applied to the assay. Both III-31-C and GSM-1 were dissolved in 100% DMSO (dimethyl sulphoxide) and diluted stepwise to ensure the same amount of DMSO in each reaction. The endogenous substrate of γ-secretase C99, carrying a C-terminal 8xHis tag and an N-terminal MBP tag, was used for another PSH cleavage assay. The concentrations of PSH and MBP-C99 in the reaction were 0.1 and 1.0 mg mL–1, respectively. The proteolytic activity of PSH is specifically suppressed by III-31-C. GSM-1 also modulates PSH similarly compared to γ-secretase in terms of the Aβ42/Aβ40 ratio (Dang et al., 2015). These results suggest that PSH can be used as a surrogate protease for screening small molecules that can regulate γ-secretase activity.

4. EXPRESSION AND PURIFICATION OF HUMAN γ-SECRETASE 4.1 Introduction to Human γ-Secretase The γ-secretase, comprising presenilin-1/2 (PS1/PS2), Nicastrin, PEN-2, and APH-1a/b, is an aspartyl intramembrane protease that catalyzes the cleavage of a wide spectrum of type I integral membrane proteins such as Notch, APP, CD44, and N-cadherin (Edbauer et al., 2003; Lammich et al., 2002; Sato et al., 2007). The γ-secretase is proposed to be the proteasome within the lipid membrane (Kopan & Ilagan, 2004). The two most thoroughly studied substrates of γ-secretase are the APP and Notch (De Strooper et al., 1999, 1998). Aberrant cleavage of APP results in production of long amyloid-β peptides that tend to form amyloid-β plaque in the brain, which is a hallmark of Alzheimer’s disease (Hardy & Higgins, 1992). The Notch signaling pathway plays a major role in cell proliferation and embryonic development, and the intracellular domain of Notch generated by γ-secretase cleavage activates transcription of specific genes. The catalytic subunit of γ-secretase is presenilin-1/2 with nine TMs. Autocatalytic activation of presenilin entails a specific cleavage between TM6 and TM7 to separate the N-terminal fragment (NTF) and the

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C-terminal fragment (CTF). PEN-2, containing three TMs, is crucial for presenilin activation and facilitates maturation of the γ-secretase (Steiner et al., 2002). Nicastrin, consisting of a large extracellular domain and a single TM, is thought to be the substrate-recruiting component. APH-1, with seven TMs, appears to stabilize the γ-secretase. Due to its crucial role in AD pathology, γ-secretase has been the target of numerous therapeutic agents that were designed to reduce the amyloid plaques. Detailed structural information of γ-secretase would greatly facilitate mechanistic understanding of its function in AD development (De Strooper, Iwatsubo, & Wolfe, 2012; Edbauer et al., 2003; St George-Hyslop & Fraser, 2012; Takasugi et al., 2003). Rigorous efforts during the past decade yielded limited structural information on γ-secretase, including low-resolution EM structure (Lazarov et al., 2006; Ogura et al., 2006; Osenkowski et al., 2009), NMR structure of the PS1 CTF (Sobhanifar et al., 2010), and crystal structure of PSH (Li, Dang, et al., 2013). Structural characterization of γ-secretase has been hindered by the technical challenge of preparing homogenous and highquality sample. After years of trials and tribulations on various expression systems, we eventually succeeded in the production of intact γ-secretase using transient expression in mammalian HEK293F cells.

4.2 Design of the Mammalian Expression Vector pMlink We designed and prepared a mammalian expression vector, named pMLink, for transient expression in HEK293F cells. The vector pMLink was modified from the pCAG plasmid. The reconstruction approach was reminiscent of pQLink that allows cloning of multiple genes in the same vector through ligation-independent cloning method (Scheich, Kummel, Soumailakakis, Heinemann, & Bussow, 2007). The procedure for the construction of the pMlink vector is summarized as follows (Fig. 4A): Step 1: Digest the original pCAG plasmid with SalI and BamHI and collect the 2000-base-pair (bp) linear fragment that contains the Ampr gene and the ColEI replication origin; Save the 2500-bp, CAG promoter containing fragment for use in step 3. Step 2: Ligate the 2000-bp fragment with a double-stranded DNA fragment that is generated by annealing two primers, each containing cutting sites for four restriction enzymes (PacI, XhoI, BglII, and SwaI). The sequences of these two primers for annealing are listed below:

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Fig. 4 Construction of pMlink vector and incorporation of the ORFs from γ-secretase. (A) The design of a mammalian expression pMlink vector for the cloning of multiple subunits of a protein complex. For details, please read the main text. (B) A working scheme for the cloning of multiple ORFs into the pMlink vector. The PacI-digested fragment can be inserted into SwaI-digested product through the ligation-independent cloning method. (C) The ORFs of PS1, APH-1, PEN-2, and nicastrin are individually cloned into the pMlink vector. Nicastrin and PEN-2 are linked together, whereas the other two are ligated. Combination of the two plasmids generates the final plasmid that contains all four ORFs of the human γ-secretase.

Forward:50 -TCGAGCTTAATTAACAACACCATTTGCTCGAGA CAGATCTGTAACAACACCATTTAAATGGAGTGGTTACAAA TGGAGTGGTTAATTAAG-30 ; Reverse:50 -GATCGCTTAATTAACCACTCCATTTGTAACCAC TCCATTTAAATGGTGTTGTTACAGATCTGTCTCGAGCAAA TGGTGTTGTTAATTAAG-30 Step 3: Digest the resulting plasmid by XhoI and BglII and ligate with the 2500-bp fragment released by SalI and BamHI digestion from step 1.

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The empty pMlink vector harbors two LINK sequences (LINK1 and LINK2) and an intervening CAG cassette, which has a promoter, a transcriptional terminator, and multiple cloning sites (Fig. 4A). The LINK1 sequence contains a PacI cutting site and the LINK2 a SwaI and a PacI sites. The PacI-digested fragment can be inserted into SwaI-digested product through the ligationindependent cloning method (Fig. 4B). Therefore, this vector allows convenient cloning of multiple ORFs for coexpression in mammalian cells. Importantly, because there is no limit to the number of ORFs, pMlink is particularly suitable for expression of protein complexes with multiple subunits. We successfully used this vector for overexpression of human γ-secretase. The ORFs of the four γ-secretase subunits, PS1, APH-1, PEN-2, and Nicastrin, are individually cloned into pMlink vector, among which PEN-2 has an N-terminal Flag tag, APH-1 carries a C-terminal HA tag, and Nicastrin contains a C-terminal V5 plus His tag. The pMlink plasmid with the largest component Nicastrin and the one with PEN-2 are linked together, whereas the plasmids for PS1 and APH-1 were also ligated. These two plasmids are combined to generate the final expression vector with all four ORFs of the human γ-secretase (Fig. 4C).

4.3 Transient Expression of Human γ-Secretase The constructed pMlink plasmid that contains the four ORFs was transformed into DH5α to obtain lipopolysaccharide-free (endo-free) plasmid for transient expression. Transfection of HEK293F cells is performed as listed: Step 1: Check the cell density until it reaches an optimal density of about 2
106 cells mL–1. Step 2: For each liter of cell culture, incubate 1.5 mg endo-free pMlink with 4 mL 25-kDa linear polyethylenimines (PEIs, 1 mg mL–1) (Polysciences; #23966; Warrington, PA) in 50 mL fresh medium for 15–30 min at room temperature. It is important to note that the optimal amounts of the plasmid and PEI should be tested for other proteins. Step 3: Add the mixture to HEK293F cells and incubate for 30 min. After culturing for about 60 h, the cells can be harvested for protein purification. Initial small-scale expression tests were examined by Western blot analysis using monoclonal antibodies against the Flag tag (ComWin; CW0287; Beijing, China), HA tag (ComWin; CW0092; Beijing, China), V5 tag (ComWin; CW0094; Beijing, China), and the NTF of PS1 (Merck Millipore; Temecula, CA). This expression strategy gives a typical yield of 0.2 mg γ-secretase for every liter of HEK293F cells.

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4.4 Purification of Human γ-Secretase 4.4.1 Required Materials 1. Reagents HEPES lysis buffer (25 mM HEPES, pH 7.4, 150 mM NaCl) Protease inhibitor cocktails, Amresco (M221; Solon, OH) Digitonin, Sigma (D141; St. Louis, MO) n-Octyl-β-D-Glucopyranoside, Anatrace (CHAPSO, C317; Maumee, OH) Anti-flag M2 affinity gel, Sigma (A220; St. Louis, MO) Flag peptide, Sigma (F3290; St. Louis, MO) Amphipol A8-35, Anatrace (A835; Maumee, OH) Bio-Beads™ SM-2 Adsorbent Media, Bio-Rad (1523920; Hercules, CA) 2. Equipment G-560E Vortex, Scientific Industries (Bohemia, NY) VCX800 Sonicator, SONICS (Newtown, CT) T10 basic ULTRA-TURRAX disperser, IKA Optima™ L-90K Ultracentrifuge, Beckman Coulter (Palo Alto, CA) Type 70 Ti rotator, Beckman Coulter (Palo Alto, CA) 50-mL PP centrifuge tubes, conical bottom with plug seal cap, Corning (China) QB-208 Multipurpose shaker, Kylin-Bell (Haimen, Jiangsu, China) Centricon, Millipore (Tullagreen, Carrigtwohill, Ireland) Size-exclusion chromatography system, GE Healthcare (Uppsala, Sweden) Superose-6 column, GE Healthcare (17-5172-01; Uppsala, Sweden) Econo-pac chromatography columns, Bio-Rad (7321010; Hercules, CA) 4.4.2 Purification of Human γ-Secretase The protocol for purification of γ-secretase is listed below (Fig. 5A): Step 1: Harvest the cells at 800
g for 10 min and resuspend the cell pellet in the HEPES lysis buffer (25 mM HEPES, pH 7.4, 150 mM NaCl) supplemented with protease inhibitor cocktails; use 30 mL lysis buffer for each liter of cells. Step 2: Disrupt the cells by sonication on ice. For every 30-mL cell resuspension, apply 50% power output for 20 cycles, each of which consists of 3 s of sonication and 3 s of pause. 3000 J is achieved after completion of the 20 cycles.

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Fig. 5 Purification of human γ-secretase and reconstitution of its enzymatic assay. (A) The flowchart is shown to describe the purification procedure for human γ-secretase. Please refer to the main text for detailed description. (B) An enzymatic activity assay for human γ-secretase (upper panel) and representative results of detection of γ-secretase cleavage products by Western blots (lower panel) (Lu et al., 2014). The assay shown in the upper panel is performed in the presence of phospholipids. The C-terminal Myc tag of AICD was detected by a monoclonal antibody. Notably, III-31-C inhibited the cleavage of C99. (C) The AlphaLISA assay is used to detect production of the Aβ peptides. Upon shared binding of acceptor and donor beads, an oxygen molecule is generated from the donor bead after photoactivation at 680 nm wavelength and transferred to the acceptor bead, which triggers an emission wavelength of 615 nm. The emission signal can be detected spectroscopically. The cleavage activities of human γ-secretase variants, each with an AD-derived mutation, were examined by the AlphaLISA assay (lower panel). The ratio of Aβ42/Aβ40 is shown for 10 variants along with the wild-type γ-secretase. Two mutations F237I and V261F led to background-level Aβ40 cleavage, disallowing the calculation of Aβ42/Aβ40 ratio. Error bars, s.d.

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Step 3: Centrifuge the disrupted cells at 150,000
g for 1 h to pellet the cell membrane. Step 4: Resuspend the cell membrane by disperser with the HEPES lysis buffer supplemented with 1% (w/v) CHAPSO and incubate at 4°C for 2 h. Step 5: Apply the resuspension to centrifugation at 150,000
g for 30 min to pellet the insoluble cell debris. Step 6: Incubate the supernatant with anti-FLAG M2 affinity gel at 4°C for 30 min. Typically, 1 mL affinity gel loaded into one column will be sufficient for 2 L of cell culture. Step 7: Wash the resin with 30 mL buffer containing 25 mM HEPES, pH 7.4, 150 mM NaCl, and 0.1% (w/v) digitonin for each column. Elute the target protein with 10 mL lysis buffer supplemented with 0.1% (w/v) digitonin and 200 μg mL–1 flag peptides. Step 8: Concentrate the elution to 12 mL with a 10-kDa cutoff centricon. Step 9: Mix the concentrated sample with amphipol A8-35 at 1:3 (w/w) and incubate at 4°C for 4 h. Step 10: Supply 60 mg Bio-Beads SM-2 to the sample and incubate overnight to eliminate detergent. Step 11: Discard the Bio-Beads after centrifugation at 14,000
g for 5 min. Apply the supernatant to gel filtration (Superose-6) to remove free amphipols. Concentrate the peak fractions to a suitable concentration for cryo-EM study.

5. ENZYMATIC ACTIVITY ASSAY FOR HUMAN γ-SECRETASE The full-length APP contains only one TM, with a large extracellular domain. APP can be cleaved by at least three proteases in vivo, namely, α-, β-, and γ-secretases. α- or β-secretase sheds the ectodomain of APP, which is a prerequisite for the second-step cleavage executed by γ-secretase. Specifically, cleavage by α-secretase results in the generation of a soluble APP-α domain in the extracellular space and an 83-residue, membrane-tethered fragment known as C83 at the carboxyl terminus (Zheng & Koo, 2011). C83 is thought to be unrelated to the development of AD. On the other hand, cleavage by β-secretase produces a soluble APP-β domain and a 99-residue, membrane-tethered fragment known as C99, which undergoes additional cleavages by γ-secretase to produce Aβ peptides (Checler, 1995; De Strooper et al., 2012; Haass & Steiner, 2002; Zheng & Koo, 2011). γ-Secretase cleaves C99 through two distinct kinds of protease activity: endoproteolysis and carboxyl terminal proteolysis (Quintero-Monzon et al.,

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2011). As an endopeptidase, γ-secretase first cleaves after the 48th or 49th residue of C99 within the transmembrane region to generate the APP intracellular domain (AICD). Then the carboxylpeptidase activity allows successive cleavage of the substrate every three or four amino acids from the C-terminus, leading to the generation of amyloid β peptides (Aβ) ranging from 37 to 49 amino acids, termed Aβ37–Aβ49 (Takami et al., 2009). AD-derived mutations in γ-secretase cause two primary consequences: dramatic reduction of the total proteolytic activity and increased molar ratio of Aβ42 over Aβ40. These consequences are a result of altered endopeptidase and carboxylpeptidase activities of γ-secretase. Intriguingly, whether these clear observations reflect the loss of function or gain of function in AD genesis remains controversial. To reconstitute the enzymatic assay, purified γ-secretase is mixed with APP-C99 that carries a Myc plus 6xHis tag at the C-terminus. The reaction buffer contains 0.5% CHAPSO, 50 mM HEPES, pH 7.4, 150 mM NaCl, 0.1% phosphatidylcholine (PC; 840053; Alabaster, AL), and 0.025% phosphatidylethanolamine (PE; 840022; Alabaster, AL). The enzyme and C99 are incubated together at 37°C for at least 3 h (Fig. 5B). Notably, in contrast to the enzymatic assays for other I-CLiPs, the phospholipids, in particular PC and PE, are indispensable for γ-secretase activity in the in vitro detergent micelle-based assay (Lu et al., 2014). For the endopeptidase activity of γ-secretase, the product AICD can be detected by Western blot using a monoclonal antibody against the C-terminal Myc tag (Fig. 5B). Incubation of γ-secretase with the substrate leads to the generation of AICD. The presenilin-specific inhibitor III-31C but not DMSO blocks the cleavage. For the carboxylpeptidase activity, three prevailing approaches have been widely utilized to probe the cleaved Aβ products. One approach relies on mass spectrometry, which provides relatively accurate measurement of Aβs of all species (Alattia et al., 2013). The second approach is to apply the cleavage products to urea SDS-PAGE to separate Aβs of different lengths for detection by specific antibodies (ChavezGutierrez et al., 2012; Klafki, Wiltfang, & Staufenbiel, 1996; QuinteroMonzon et al., 2011; Wiltfang et al., 2002). The third approach is AlphaLISA (Perkin Elmer; Boston, MA), which is somewhat similar to the classic ELISA assay (Veugelen, Saito, Saido, Chavez-Gutierrez, & De Strooper, 2016). Two types of beads, acceptor and donor, are each attached to a monoclonal antibody. The monoclonal antibody attached to the acceptor beads recognizes a specific region in Aβ40 or Aβ42; in contrast, the antibody attached to the donor beads recognizes a common region in Aβ

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peptides. Upon shared binding of a specific Aβ peptide, an oxygen molecule is generated through photoactivation of the donor bead at 680 nm wavelength and transferred to the acceptor bead, which has an emission wavelength of 615 nm. The emission signal can be detected spectroscopically (Fig. 5C). The detection limit of the AlphaLISA assay for Aβ40 and Aβ42 is approximately 30 pg mL–1 or about 7 pM, which allows successful detection of Aβ production at a very low quantity (Bai et al., 2015). The standard protocol of the AlphaLISA assay is as follows: Step 1: Generate standard curves with lyophilized Aβ40 and Aβ42 peptides to quantify specific Aβs. Step 2: Add 8 μL AlphaLISA Aβ1-40/1-42 acceptor beads into 2 μL reaction sample and incubate at 22°C for 1 h. Step 3: Add 10 μL AlphaLISA donor beads and incubate in the dark at 22°C for another 30 min. Step 4: Record the spectroscopic reading of the samples by EnvisionAlpha Reader (PerkinElmer; Boston, MA). Step 5: Calculate the amount of Aβ peptides by comparing with the standard curves. Based on the assay system, disease-derived PS1 mutants exhibit distinct Aβ40 and Aβ42 production compared to wild-type γ-secretase (Fig. 5C).

6. STRUCTURES OF I-CLiPs The methods described earlier facilitated structural determination of GlpG, mjS2P, PSH, and human γ-secretase. In our case, the transmembrane core domain of GlpG, representing the core part of the rhomboid serine protease, forms a pseudo dimer in one asymmetric unit (Wu et al., 2006). It contains six TMs, with catalytic residues Ser201 and His254 located in TM4 and TM6, respectively, in a V-shaped cavity that opens to the extracellular side. Moreover, the imidazole side chain of His254 forms a hydrogen bond with the hydroxyl group of Ser201, which appears to be ready to initiate a nucleophilic attack toward the substrate. The N-terminal portion of GlpG, including TM1-3 and Loop1, may serve as the structural scaffold, whereas the C-terminal half of the molecule, comprising TM4-6, constitutes a functional entity (Fig. 6A). Several structures of GlpG in different conformations have been reported, some of which were obtained in the presence of inhibitors (Ben-Shem et al., 2007; Brooks, Lazareno-Saez, Lamoureux, Mak, & Lemieux, 2011; Cho, Dickey, & Urban, 2016;

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Fig. 6 Structures of I-CLiPs. (A) Crystal structure of GlpG (Wu et al., 2006). The catalytic residues Ser201 and His254 are colored yellow. TM5 (orange) serves as a putative lateral gate. (B) Crystal structure of the S2P homolog from Methanocaldococcus jannaschii (mjS2P) (Feng et al., 2007). The catalytic zinc ion (gray) is coordinated by two histidine residues (His54 and His58) from the HExxH motif and an aspartate residue (Asp148) on TM2. TM1, TM5, and TM6 may represent the mobile gate domain, whereas the TM2–4 serves as a conserved core domain. (C) Crystal structure of PSH, the presenilin-1/SPP homolog from Methanoculleus marisnigri JR1 (Li, Dang, et al., 2013). The two catalytic aspartate residues Asp162 and Asp220 (yellow) are located in TM6 and TM7. (D) Atomic structure of human γ-secretase is shown in cartoon representation (Bai et al., 2015). N-linked glycans are displayed in sticks. (E) Structural alignment between PS1 and PSH. (F) A close-up view on the catalytic residues and the PAL motifs of PS1 and PSH. The PDB codes are 2NRF (GlpG), 3B4R (mjS2P), 4HYG (PSH), and 5A63 (γ-secretase).

Lemieux et al., 2007; Vinothkumar, 2011; Vinothkumar et al., 2010; Wang & Ha, 2007; Wang et al., 2006; Xue & Ha, 2012; Zoll et al., 2014). The structural prototype of intramembrane metalloprotease is the crystal structure of site-2 protease (S2P) from M. jannaschii. The catalytic zinc ion is coordinated by two histidine residues (His54 and His58) from the HExxH motif and an aspartate residue (Asp148) on TM2 (Fig. 6B). Similar to GlpG, the core domain of mjS2P also consists of six TMs, which display two different conformations—close and open—in one asymmetric unit. Comparison between the two conformations shows that TM1, TM5, and TM6 may represent the mobile gate domain, whereas the TM2-4 serves as a conserved core domain (Feng et al., 2007).

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The archaeal homolog of presenilin/SPP PSH from Methanoculleus marisnigri JR1 offers the first glimpse of structural insights into aspartyl intramembrane protease. PSH displays a novel membrane protein fold with nine TMs. The NTD, comprising TM1–6, forms a horseshoe-shaped structure around the CTD of TM7–9 (Fig. 6C). The two catalytic aspartate res˚, idues are located in TM6 and TM7 and separated by a distance of 6.7 A which is larger than that in an activated aspartate protease. Therefore, conformation of PSH in the crystals may represent that of an inactive protease (Li, Dang, et al., 2013). The structure of human γ-secretase contains a soluble domain of Nicastrin and 20 TMs organized into a horseshoe appearance. The nine TMs of PS1 are placed between APH-1 and PEN-2. The single TM of Nicastrin is located besides APH-1 at the edge of the thick end of the transmembrane region (Fig. 6D). The catalytic residues, Asp257 on TM6 and Asp385 on TM7, are arranged on the convex side of the horseshoe. The distance between the Cα atoms of Asp257 and Asp385 measures approximately 10.6 A˚, considerably longer than that in an activated aspartyl protease. The atomic structure of γ-secretase revealed detailed interactions among the four components and lipids (Bai et al., 2015). Comparison between PS1 and PSH demonstrates a similar overall conformation and conserved locations of the two catalytic aspartic acids and the putative substrate-binding PAL motifs (Fig. 6E and F).

7. CONCLUDING REMARKS Structure determination of I-CLiPs allows mechanistic understanding of how peptide bond hydrolysis occurs in the highly hydrophobic membranous environment. A prerequisite for structural investigation is a successful preparation of homogenous sample with reasonable solution behavior. Unfortunately, this usually represents a bottleneck, owing to the poor expression level of an I-CLiP and its unruly solution behavior. Acquisition of a well-behaved I-CLiP for structural and biochemical study requires homolog screening and protein engineering. The successes of dealing with the I-CLiPs have offered useful tips for working with other classes of membrane proteins. Natural variants and homologs from a large number of representative species provide a database of potential target proteins, from which a construct amenable for crystallization may be chosen through extensive expression and purification screening. Sequence-based protein engineering may improve the solution behavior of membrane proteins; one of the most remarkable examples of this method is the

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identification of PSH variants that eventually gave rise to diffraction-quality crystals. In addition, limited proteolysis proves to be an important approach for the identification of stable core domain and the removal of flexible surface loops. The stable conformation of a core domain greatly improves the chances of crystallization. Moreover, the choice and concentrations of detergents play an essential role in the preparation of suitable sample for structural and biochemical characterization. Typically, a screening process involving multiple types of detergents is required. Furthermore, as suggested by the experience with γ-secretase, eukaryotic membrane proteins are frequently misfolded or aggregated when expressed in bacteria; hence eukaryotic expression system, especially mammalian cells, will be of increasing significance in the future. Structural biology is experiencing an unprecedented wave of revolution by single particle cryo-EM, largely because of the development and application of direct electron detector (Bai, Fernandez, McMullan, & Scheres, 2013; Li, mooney et al., 2013; Liao, Cao, Julius, & Cheng, 2013, 2014). The biggest hurdle in traditional, X-ray-based structural investigation of macromolcules is to obtain well-ordered, diffracting crystals. This task is completely eliminated in cryo-EM analysis. More importantly, compositional and conformational homogeneity, which is usually required for X-ray crystallography, is no longer a prerequisite for cryo-EM. The different conformations can be differentiated by two-dimensional and threedimensional classifications during cryo-EM image analysis. The primary challenges for cryo-EM are purification of good-quality sample and preparation of sample grids for imaging. Nonetheless, the high-throughput aspect and mature methodology of X-ray crystallography cannot yet be paralleled by cryo-EM analysis. Thus, in the foreseeable future, X-ray crystallography will continue to dominate structure determination of proteins with molecular weight of 100-kDa or less. The available structural information on I-CLiPs has revealed significant insights into mechanistic understanding of regulated intramembrane proteolysis. Despite these advances, many fundamental questions remain to be addressed. For instance, how is the substrate recognized by the I-CLiPs and delivered to the active site? The conformation of an I-CLiP must be highly dynamic during each cycle of substrate binding and hydrolysis. Do the conformations of an I-CLiP represent a toggle switch, several distinct states, or a continuum between two extreme states? How can these distinct conformations of an I-CLiP be captured and validated? For peptide bond hydrolysis, the transmembrane helix of the substrate protein needs to be

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presented in an extended conformation. Does this occur simply by thermal motion or is it facilitated by the I-CLiP? Answers to these questions await future investigations by structural biology and molecular biophysics. This chapter mainly discusses the expression and purification of the I-CLiPs for which we have hands-on experience, along with the development of proteolytic activity assays for these I-CLiPs. The methods described here may be adopted to other intramembrane proteases and enzymes.

ACKNOWLEDGMENTS We apologize to those colleagues whose important contributions are not cited in this chapter due to space limitations. This work was supported by funds from the Ministry of Science and Technology and the National Natural Science Foundation of China. The authors declare no competing financial interests. Correspondence and requests for materials should be addressed to R.Z. ([email protected]), G.Y. ([email protected]), or Y.S. ([email protected]).

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CHAPTER SIX

Analyzing Amyloid-β Peptide Modulation Profiles and Binding Sites of γ-Secretase Modulators J. Trambauer*, A. Fukumori*,†,1, B. Kretner*,†,2, H. Steiner*,†,3 *Biomedical Center (BMC), Metabolic Biochemistry, Ludwig-Maximilians-University Munich, Munich, Germany † German Center for Neurodegenerative Diseases (DZNE), Munich, Germany 3 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Methods 2.1 Assaying GSM Activity 2.2 Target Identification of GSMs Using Photoaffinity Labeling Acknowledgments References

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Abstract γ-Secretase is a key player in the pathogenesis of Alzheimer’s disease (AD). The intramembrane-cleaving enzyme initially cleaves a C-terminal fragment of the amyloid precursor protein (APP) at the ε-site within its transmembrane domain to release the APP intracellular domain. Subsequent stepwise carboxy-terminal trimming cleavages eventually release amyloid-β (Aβ) peptides of 37–43 amino acids into the extracellular space. Aβ42 as well as the much less abundant Aβ43 species are highly aggregation prone and can deposit as plaques in the brains of affected patients, which are widely believed to be causative of AD. Disappointingly, due to lack of efficacy and side effects likely attributable to the inhibition of the crucial substrate Notch, inhibitors of γ-secretase that lower Aβ generation failed in clinical trials of AD. There is hope, however, that recently developed potent γ-secretase modulators (GSMs) provide a safer approach for disease modification. These compounds have the unique property of primarily shifting the generation of Aβ42 toward that of shorter peptides without affecting the ε-site cleavage of Notch and other substrates. In this chapter, we describe methods to investigate how GSMs affect the activity of the enzyme as well as how their molecular targets are identified. 1 2

Present address: Department of Psychiatry, Health Care Center, Osaka University, Toyonaka, Japan. Present address: Division for Neurodegenerative Diseases, Department of Neurology, Technical University Dresden, Germany.

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1. INTRODUCTION γ-Secretase is a pivotal protease essential for life and the most complicated intramembrane-cleaving enzyme known (De Strooper, Iwatsubo, & Wolfe, 2012; Sun, Li, & Shi, 2016). The enzyme is an integral membrane protein complex composed of the four subunits presenilin (PS), which harbors the active site, nicastrin (NCT), APH-1, and PEN-2 (De Strooper et al., 2012). γ-Secretase cleaves numerous type I membrane proteins as substrates (Haapasalo & Kovacs, 2011), whose cleavage requires the presence of short ectodomains (Struhl & Adachi, 2000), usually obtained by shedding events. Cleavage of its major substrate Notch, releases the Notch intracellular domain, a signaling molecule that is not only essential for cell differentiation in development but also critical later in life (Kopan & Ilagan, 2009). Due to its involvement in the pathogenesis of Alzheimer’s disease (AD) by producing the likely disease-causing amyloid-β (Aβ) peptides, γ-secretase is a key AD drug target and consequently has been the most intensively studied intramembrane protease to date (De Strooper et al., 2012). Aβ is a heterogeneous mixture of small peptides 37–43 amino acids in length that are generated by sequential cleavage of the amyloid precursor protein (APP) by β- and γ-secretase (Lichtenthaler, Haass, & Steiner, 2011). Following β-secretase cleavage, which removes the bulk of the APP ectodomain, γ-secretase processes the resultant APP C-terminal fragment, termed APP CTFβ or C99. This cleavage is very complex and involves both endopeptidase and carboxypeptidase activities of the enzyme (Morishima-Kawashima, 2014). After an initial endopeptidase cleavage at the ε-site close to the cytoplasmic transmembrane domain border, which releases the APP intracellular domain, the resultant long Aβ is processed to shorter Aβ peptides. Depending on the initial ε-site attack, two principal Aβ product lines are initiated (Qi-Takahara et al., 2005; Takami et al., 2009). In one product line, the major species Aβ40 is generated in a stepwisemanner from Aβ49, Aβ46, and Aβ43. A second product line starting from Aβ48 generates Aβ45 and then Aβ42. Both lines also give rise to small amounts of the shorter peptides Aβ37 that is generated from the Aβ40 line and Aβ38 that is generated from the Aβ42 line. The product lines can also be crossed (Okochi et al., 2013). The longer Aβ species Aβ42 and Aβ43 are highly aggregation-prone neurotoxic peptides that are believed to trigger a cascade of pathological alterations in the brain that ultimately lead to neurodegeneration and dementia (Selkoe & Hardy, 2016). This “amyloid cascade” hypothesis is strongly supported by the fact that the large majority

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of the clinical mutations associated with an early onset familial AD (FAD) locate in the PS1 and PS2 catalytic subunits of γ-secretase as well as around the γ-secretase cleavage sites of the APP transmembrane domain and cause an increase in the ratios of Aβ42 or Aβ43 to Aβ40 (Weggen & Beher, 2012). The failure of γ-secretase inhibitors (GSIs) in large clinical trials of AD has led to a big disappointment in the field (De Strooper & Chavez Gutierrez, 2015). However, suboptimal drug dosing schemes as well as no or too little APP substrate specificity of the compounds tested so far can well explain the failure of these compounds as well as their association with severe Notchrelated side effects. The failure of GSIs in clinical trials shifts the focus to likely more safer and superior drugs, the so-called γ-secretase modulators (GSMs). These compounds show preferential lowering of Aβ42 without affecting ε-site cleavage of γ-secretase substrates (Weggen et al., 2001, 2003). Concomitantly to lowering Aβ42, these compounds also increase the generation of short Aβ species, most commonly Aβ38, i.e., they alter γ-secretase cleavage specificity (Weggen et al., 2001). While initially certain nonsteroidal antiinflammatory drugs (NSAIDs) were identified as GSMs with micromolar potencies (Weggen et al., 2001), a number of structurally different second-generation compounds with better potencies and IC50 values in the low nanomolar range have been identified in drug-screening programs of pharmaceutical companies and are further developed into clinical candidates (Bursavich, Harrison, & Blain, 2016; Crump, Johnson, & Li, 2013). Based on their chemical structures, GSMs can be divided into two main structural classes. The so-called acidic GSMs all contain a carboxyl group, which is essential for their Aβ42-lowering activity. Modification of the carboxyl group reverses the activity of the GSM to become an inverse modulator, which increases long Aβ species while lowering the short Aβ species (Narlawar, Baumann, Czech, & Schmidt, 2007; Ohki et al., 2011). Certain NSAIDs such as sulindac sulfide (Weggen et al., 2001) belong to this class as well as the NSAIDbased GSM-1 (Page et al., 2008), which is a prototype of the more potent second-generation acidic GSMs. The other major structural class is that of the so-called nonacidic or bridged aromatic GSMs, which are structurally composed of four linearly aligned aromatic building blocks. These include compounds initially identified by Neurogenetics (Kounnas et al., 2010) and many derivatives thereof such as RO-02 developed by Roche (Ebke et al., 2011). Examples of acidic and bridged aromatic GSMs are shown in Fig. 1. More recently, endogenous steroids (Jung et al., 2013) and other triterpene compounds identified by Satori (Findeis et al., 2012) have been reported to act as GSMs. Unlike the low potency NSAIDs (Czirr et al., 2007; Page et al., 2008), most second-generation GSMs are capable of lowering the increased levels

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Fig. 1 Structures of acidic (GSM-1) and bridged aromatic (RO-02) secondgeneration GSMs.

of Aβ42 generated by PS FAD mutants (Kretner et al., 2011; Szaruga et al., 2015). Only few mutants such as the aggressive PS1 L166P mutant are resistant to the Aβ42-lowering activity of these GSMs (Kretner et al., 2011). Here, we provide detailed protocols how the modulatory activity of GSMs can be studied using cell-based models and how their molecular targets can be identified using photoactivatable cross-linkable GSM derivatives.

2. METHODS 2.1 Assaying GSM Activity Acidic and bridged aromatic GSMs modulate the generation of Aβ species differently. Acidic GSMs lower Aβ42 and increase Aβ38 (Page et al., 2008; Weggen et al., 2001). In contrast, bridged aromatic compounds typically lower both Aβ42 and Aβ40 while increasing Aβ38 and Aβ37 (Kounnas et al., 2010). Yet another profile has been reported for the Satori triterpenes, which lower Aβ42 and Aβ38, while leaving Aβ40 unaffected and increasing Aβ39 and Aβ37 (Findeis et al., 2012). The modulatory activity of GSMs can be assessed conveniently in cellbased assays. For ease of analysis, cell lines such as HEK293 cells stably overexpressing Swedish mutant APP (HEK293/sw), which produce high amounts of Aβ (Citron et al., 1992), are frequently used to facilitate analysis of Aβ species. To analyze the efficacy of GSMs on mutant γ-secretases, we use HEK293/sw cells stably transfected with wt control or mutant PS constructs (Kretner et al., 2011; Page et al., 2008). In the following sections, we describe two different experimental setups to assess GSM activity. Simple drug treatment experiments are used to

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analyze overall GSM activity and GSM dose–response experiments are used for potency determination that give more detailed insight into the critical parameters of modulation such as values for the inhibitor concentration causing 50% inhibition (IC50) of Aβ42 generation. 2.1.1 Cell Culture and Drug Treatments Equipment: Heating block Microcentrifuge tubes (1.5 mL) Micropipette Micropipette tips Refrigerated benchtop centrifuge (1.5 mL tubes) Materials: Dulbecco’s Modified Eagle Medium, high glucose, GlutaMAX™ (Thermo Scientific, Waltham, MA, USA; #61965026) (DMEM) Dimethyl sulfoxide (DMSO) Fetal calf serum (FCS) Poly-L-lysine Solutions: DMEM/FCS DMEM Containing 10% FCS Component

Final Concentration

Stock

Amount

FCS

10%

—

50 mL

DMEM

—

—

Add to 500 mL

Protocol: 1. Preparation of conditioned medium of GSM-treated cells a. Seed HEK293/sw cells in DMEM/FCS on poly-L-lysine coated 6-well plates. Grow them to
80% confluence. b. Prepare at least one dilution of the GSM in DMEM/FCS together with a DMSO containing vehicle control. GSM concentrations chosen are typically around the IC50 for Aβ42 generation and/or 5- to 10-fold higher. Make sure that the DMSO concentration in control and GSM samples does not exceed 0.1%. c. Remove the medium from the cells, add 1 mL of the prepared dilutions, and incubate the cells for 16–18 h. 2. Preparation of conditioned medium for dose–response experiments a. Seed HEK293/sw cells in DMEM/FCS on poly-L-lysine coated 24-well plates. Grow them to
80% confluence.

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b. Prepare a serial dilution of the GSM in DMEM/FCS (triplicates of seven concentrations) typically ranging from 1010 to 104 M and DMSO as control. Depending on GSM potency, the concentration range may be adjusted. c. Remove the medium from the cells, add 500 μL of the prepared GSM dilutions, and incubate the cells for 16–18 h. 3. Collect the conditioned medium and centrifuge (10 min, 4°C, 10,000  g) to clear cellular debris. Use the cleared medium either directly for MSD sandwich immunoassay or for immunoprecipitation of Aβ followed by Tris–Bicine urea SDS-PAGE or matrix-assisted laser desorption/ionization (MALDI)-time of flight (TOF) mass spectrometry (MS) to analyze the complete profile of Aβ species or store it until further use at 20°C.

2.1.2 MSD Sandwich Immunoassay Analysis of Aβ Species Equipment: Heating block Microcentrifuge tubes Micropipette, multichannel pipette Micropipette tips Meso Scale Discovery (MSD, Rockville, MD, USA) multiplex reader (e.g., MESO™ QuickPlex SQ 120) Plate shaker (at RT and 4°C) Refrigerated benchtop centrifuge (1.5 mL tubes) Materials: Aβ peptide standard solutions Bovine serum albumin (BSA) Disodium hydrogen phosphate (Na2HPO4) Dulbecco’s Modified Eagle Medium, high glucose, GlutaMAX™ (Thermo Scientific, Waltham, MA, USA; #61965026) (DMEM) FCS MSD Streptavidin Gold Multiarray® 96-well plates Potassium chloride (KCl) Potassium dihydrogen phosphate (KH2PO4) Sodium chloride (NaCl) Tween 20 Antibodies: Capture antibody: Antibody recognizing all Aβ species such as 6E10 (Covance, Princeton, NJ, USA; SIG-39320) or, as we use, 2D8

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(Shirotani, Tomioka, Kremmer, Haass, & Steiner, 2007). The capture antibody is biotinylated using standard procedures. Detection antibodies: MSD-GOLD-SULFO-tagged detection antibodies specific to the C-termini of Aβ38, Aβ40, and Aβ42 (MSD, Rockville, MD, USA). Other suitable end-specific anti-Aβ antibodies such as BAP24 (anti-Aβ40) and BAP15 (anti-Aβ42) (Brockhaus et al., 1998; Czech et al., 2007), as used by our laboratory, can be SULFO-tagged using the manufacturer’s protocol. Solutions: 10 × PBS-T 0.5% Tween 20 in 10 × PBS, pH 7.4 Component

Final Concentration

Stock

Amount

NaCl

1370 mM

—

80 g

KCl

27 mM

—

2g

Na2HPO4

100 mM

—

14.2 g

KH2PO4

18 mM

—

2.5 g

Tween 20

0.5%

—

5 mL

H2 O

—

—

Add to 1 L

Adjust pH to 7.4 before adding H2O to 1 L

Blocking Buffer 1% BSA, 0.05% Tween 20 in PBS, pH 7.4 Component

Final Concentration

Stock

Amount

PBS-T

1

10 

50 mL

BSA

1%

—

5g

H2O

—

—

Add to 500 mL

Adjust pH to 7.4 after addition of BSA

Washing Buffer 0.05% Tween 20 in PBS, pH 7.4 Component

Final Concentration

Stock

Amount

PBS-T

1

10 

50 mL

H2O

—

—

Add to 500 mL

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DMEM/FCS DMEM Including 10% FCS Component

Final Concentration

Stock

Amount

FCS

10%

—

50 mL

DMEM

—

—

Add to 500 mL

Protocol: 1. Add 100 μL blocking buffer per well on a MSD Streptavidin Gold Multiarray® 96-well plate and shake for 30 min at RT. 2. Dilute biotin-labeled anti-Aβ capture antibody to an appropriate concentration (e.g., 2.4 μg/mL for biotinylated 2D8) in blocking buffer. Add 100 μL per well and shake for 60 min at RT, or overnight at 4°C to coat the plate. 3. To wash the plate, shake two times for 5 min with washing buffer at RT. Remove liquid completely between the steps by emptying the 96-well plate onto a tissue. 4. Prepare Aβ standards in DMEM/FCS. Make a serial dilution using 30,000 pg/mL as highest Aβ concentration and dilute peptide standards six times 1:3 with DMEM/FCS. Load 75 μL per well. 5. Load 75 μL sample (see Section 2.1.1) on the plate. Depending on the Aβ amount, samples may have to be diluted in DMEM/FCS. 6. Add 25 μL of MSD-GOLD-SULFO-tagged Aβ-species specific detection antibodies. Dilute the antibodies in blocking buffer (usually 1:500–1:2000) depending on antibody sensitivity and Aβ amounts in the sample. 7. Shake for 2 h at RT in the dark (e.g., by covering plate with aluminum foil). 8. Wash three times for 5 min with washing buffer. Shield the plate from light while shaking. 9. Add 100 μL 1  reading buffer and measure immediately on a MSD multiplex reader. Analysis: The measured concentrations of the Aβ species can be analyzed in two different ways. I. The concentrations of the different species can be divided through the total amount of Aβ measured (i.e., the sum of Aβ38, Aβ40, and Aβ42) to calculate Aβ ratios, which are then plotted in comparison to those of nontreated cells. II. The concentrations of the different Aβ species of the GSM-treated cells are normalized to those of the vehicle treated cells, which are set to 100%. Note that when using this method, plots showing only

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single Aβ species can be misleading, as changes in their concentration may result from a change in total Aβ production rather than modulation of the production of the respective species. To analyze dose–response experiments, data values are calculated as in step II and plotted against the logarithm of the GSM concentration and expressed as percentage of vehicle treated (see Fig. 2A). The graph

Fig. 2 Modulation of γ-secretase activity by acidic and nonacidic GSMs. (A) Dose– response curves of GSM-1a and RO-02b for Aβ38, Aβ40, and Aβ42 species measured by sandwich immunoassay. (B) Tris–Bicine urea SDS-PAGE analysis of Aβ species. (C) MALDI-TOF MS analysis of Aβ species.a In (B) and (C), HEK293/sw cells stably overexpressing wt PS1 were used for analysis. a

b

This research was originally published in The Journal of Biological Chemistry. Kretner, B., Fukumori, A., Gutsmiedl, A., Page, R. M., Luebbers, T., Galley, G., Baumann, K., Haass, C., & Steiner, H. (2011). Attenuated Aβ42 responses to low potency γ-secretase modulators can be overcome for many pathogenic presenilin mutants by second-generation compounds. The Journal of Biological Chemistry, 286, 15240–15251. © the American Society for Biochemistry and Molecular Biology. This research was originally published in The Journal of Biological Chemistry. Ebke, A., Luebbers, T., Fukumori, A., Shirotani, K., Haass, C., Baumann, K., & Steiner, H. (2011). Novel γ-secretase enzyme modulators directly target presenilin protein. The Journal of Biological Chemistry, 286, 37181–37186. © the American Society for Biochemistry and Molecular Biology.

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is plotted using nonlinear regression by an appropriate data analysis program (e.g., GraphPad Prism). The concentration at the inflection point represents the IC50 of the GSM for the respective species (Aβ40 or Aβ42). As the MSD sandwich immunoassay is limited to the analysis of certain Aβ species by the availability of end-specific Aβ antibodies (typically antibodies against Aβ38, Aβ40, and Aβ42), additional methods are needed to assess complete Aβ profiles. To this end, separation of Aβ species by Tris–Bicine urea SDS-PAGE (Fig. 2B) as well as MALDI-TOF MS analysis (Fig. 2C) is used. These procedures are described in the following sections.

2.1.3 Tris–Bicine Urea SDS-PAGE Analysis of Aβ Species Equipment: Centrifuge tubes (15 and 50 mL) Heating block Microcentrifuge tubes (1.5 mL) Micropipette, multichannel pipette Micropipette tips Polyacrylamide gel electrophoresis and immunoblotting equipment Refrigerated benchtop centrifuge (1.5 mL tubes) Materials: Acrylamide (40% Acrylamide/Bis-Acrylamide 19:1, 5% Crosslinker) Bis–Tris Bicine Bromophenol blue Concentrated (96%) sulfuric acid (H2SO4) Disodium hydrogen phosphate (Na2HPO4) DMSO Ethylenediaminetetraacetic acid (EDTA) Hydrochloric acid (HCl) I-Block™ (Tropix®, Applied Biosystems (Thermo Scientific, Waltham, MA, USA; T2015) β-Mercaptoethanol Milli-Q water (H2O) NP-40 Potassium chloride (KCl) Potassium dihydrogen phosphate (KH2PO4)

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Protein A-Sepharose (PAS) Protein G-Sepharose (PGS) Saccharose Sodium chloride (NaCl) Sodium dodecylsulfate (SDS) Sodium hydroxide (NaOH) Tris Tween 20 Urea Antibodies: Immunoprecipitation: Antibodies that immunoprecipitate all Aβ species. We use antibody 3552 (Yamasaki et al., 2006). Immunoblotting: Antibodies that detect all Aβ species, e.g., 6E10 (Covance, Princeton, NJ, USA; SIG-39320). We use antibody 2D8 (Shirotani et al., 2007). Solutions: 4 × Separation Gel Buffer 1600 mM Tris, 400 mM H2SO4 Component

Final Concentration

Stock

Amount

Tris

1600 mM

—

48.46 g

H2SO4

400 mM

—

5.55 mL

H2O

—

—

Add to 250 mL

2 × Spacer Gel Buffer 800 mM Bis–Tris, 200 mM H2SO4 Component

Final Concentration

Stock

Amount

Bis–Tris

800 mM

—

16.74 g

H2SO4

200 mM

—

1.11 mL

H2O

—

—

Add to 100 mL

2 × Stacking Gel Buffer 720 mM Bis–Tris, 320 mM Bicine Component

Final Concentration

Stock

Amount

Bis–Tris

720 mM

—

37.66 g

Bicine

320 mM

—

13.05 g

H2O

—

—

Add to 250 mL

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2 × Tris–Bicine Urea Sample Buffer 720 mM Bis–Tris, 320 mM Bicine, 2% SDS, 5% β-Mercaptoethanol, 30% Saccharose, Bromophenol Blue Component

Final Concentration

Stock

Amount

Bis–Tris

720 mM

—

4.52 g

Bicine

320 mM

—

1.57 g

SDS

2%

20%

3 mL

β-Mercaptoethanol

5%

—

1.5 mL

Saccharose

30%

—

9g

H2O

—

—

Add to 30 mL

Add a spatula tip of bromophenol blue to the prepared sample buffer 1.4 × Tris–Bicine Urea sample buffer 504 mM Bis–Tris, 224 mM Bicine, 1.4% SDS, 3.5% β-mercaptoethanol, 21% saccharose, bromophenol blue

To obtain a 1.4  Tris–Bicine urea sample buffer, prepare 2  buffer first, then dilute to 1.4  STEN-NaCl 50 mM Tris–HCl, pH 7.6, 500 mM NaCl, 2 mM EDTA, 0.2% NP-40 Component

Final Concentration

Stock

Amount

Tris–HCl, pH 7.6

50 mM

1M

25 mL

NaCl

500 mM

5M

50 mL

EDTA, pH 8

2 mM

0.5 M

2 mL

NP-40

0.2%

—

1 mL

H2O

—

—

Add to 500 mL

STEN-SDS 50 mM Tris–HCl, pH 7.6, 150 mM NaCl, 2 mM EDTA, 0.2% NP-40, 0.1% SDS Component

Final Concentration

Stock

Amount

Tris–HCl, pH 7.6

50 mM

1M

25 mL

NaCl

150 mM

5M

15 mL

SDS

0.1%

20%

2.5 mL

EDTA, pH 8

2 mM

0.5 M

2 mL

NP-40

0.2%

—

1 mL

H2O

—

—

Add to 500 mL

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STEN 50 mM Tris–HCl, pH 7.6, 150 mM NaCl, 2 mM EDTA, 0.2% NP-40 Component

Final Concentration

Stock

Amount

Tris–HCl, pH 7.6

50 mM

1M

25 mL

NaCl

150 mM

5M

15 mL

EDTA, pH 8

2 mM

0.5 M

2 mL

NP-40

0.2%

—

1 mL

H2O

—

—

Add to 500 mL

Anode Buffer 200 mM Tris, 50 mM H2SO4 Component

Final Concentration

Stock

Amount

Tris

200 mM

—

121.14 g

H2SO4

50 mM

—

13.9 mL

H2 O

—

—

Add to 5 L

Cathode Buffer 200 mM Bicine, 0.25% SDS, 100 mM NaOH Component

Final Concentration

Stock

Amount

Bicine

200 mM

—

65.3 g

SDS

0.25%

20%

25 mL

NaOH

100 mM

—

8g

H2 O

—

—

Add to 2 L

PBS 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4 Component

Final Concentration

Stock

Amount

NaCl

137 mM

—

8g

KCl

2.7 mM

—

0.2 g

Na2HPO4

10 mM

—

1.42 g

KH2PO4

1.8 mM

—

0.25 g

H2O

—

—

Add to 1 L

Adjust pH to 7.4 before adding H2O to 1 L

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10 × PBS-T 0.5% Tween 20 in 10 × PBS, pH 7.4 Component

Final Concentration

Stock

Amount

NaCl

1370 mM

—

80 g

KCl

27 mM

—

2g

Na2HPO4

100 mM

—

14.2 g

KH2PO4

18 mM

—

2.5 g

Tween 20

0.5%

—

5 mL

H2O

—

—

Add to 1 L

Adjust pH to 7.4 before adding H2O to 1 L 0.2% I-Block 0.2% I-Block™ in 0.05% Tween 20/PBS, pH 7.4 Component

Final Concentration

Stock

Amount

PBS-T

1

10 

50 mL

I-Block™

0.2%

—

1g

H2O

—

—

Add to 500 mL

Gel solutions: The following amounts are calculated for one gel (9.6 cm high, 10.1 cm wide, and 1.5 mm thick) using the Hoefer Mighty Small II® system. 11% Separation Gel 400 mM Tris, 100 mM H2SO4, 0.1% SDS, 8 M Urea, 11% Acrylamide Component

Final Concentration

Stock

Amount

Separation gel buffer

1

4

2.5 mL

H2O

—

—

1 mL

Urea

8M

—

4.8 g

SDS

0.1%

20%

50 μL

Acrylamide

11%

40%

2.75 mL

APS

—

10%

40 μL

TEMED

—

—

5 μL

—

—

Σ 10 mL

Dissolve urea directly in buffer + H2O by slight heating before addition of SDS and acrylamide

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6% Spacer Gel 100 mM H2SO4, 400 mM Bis–Tris, 0.1% SDS, 6% Acrylamide Component

Final Concentration

Stock

Amount

Spacer gel buffer

1

2

2 mL

H2 O

—

—

1360 μL

SDS

0.1%

20%

20 μL

Acrylamide

6%

40%

600 μL

APS

—

10%

16 μL

TEMED

—

—

4 μL

—

—

Σ 4 mL

9% Stacking Gel 360 mM Bis–Tris, 160 mM Bicine, 0.1% SDS, 9% Acrylamide Component

Final Concentration

Stock

Amount

Stacking gel buffer

1

2

1.5 mL

H2O

—

—

740 μL

SDS

0.1%

20%

15 μL

Acrylamide

9%

40%

675 μL

APS

—

10%

18 μL

TEMED

—

—

6 μL

—

—

Σ 6 mL

Protocol: 1. Prepare the gel solutions. 2. Cast the gel: The separation gel should have a length of 7 cm, stacking, and spacer gels 1.3 cm each. 3. Prepare the samples using 2  Tris–Bicine urea sample buffer. For Aβ detection from medium, one usually has to concentrate the sample by immunoprecipitation. Samples with higher Aβ levels such as brain lysates of APP transgenic mice (Brendel et al., 2015; Page et al., 2008) and in vitro γ-secretase cleavage assays (Ebke et al., 2011; Winkler et al., 2009) can be loaded directly on the gel. 4. Immunoprecipitation of Aβ: a. Take the conditioned medium (usually 1 mL, see Section 2.1.1). Add 20 μL PAS or PGS beads depending on host species in which antibody was raised and/or IgG subclass and an appropriate amount

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anti-Aβ antibody. For a rabbit polyclonal serum antiserum (e.g., antibody 3552), we use 2 μL antibody/20 μL beads. b. Shake for 60 min at RT or overnight at 4°C. c. Collect the beads by centrifugation (1 min, 4°C, 2500  g), and carefully aspirate the supernatant. d. Wash the beads subsequently with STEN-NaCl, STEN-SDS, and STEN buffer. Add 1 mL of buffer, vortex briefly, centrifuge (1 min, 4°C, 2500  g) again, and carefully aspirate the supernatant. After the third wash, carefully remove the complete supernatant from the beads. e. Add 15 μL 1.4  Tris–Bicine urea sample buffer, and incubate 5 min at 95°C. 5. Fill gel chambers with anode and cathode buffer. Start the gel run with 80 V and increase voltage to 120 V after 20 min. Keep a current of 30 mA per gel by adjusting the voltage to up to 160 V during the remaining run. 6. Blot electrophoresed proteins onto nitrocellulose membrane. After the transfer, boil the membrane for 5 min in PBS. 7. Shake for 1 h at RT in 0.2% I-Block to block the membrane. 8. Use antibodies recognizing all Aβ species for detection. Tip: Load synthetic Aβ peptide standards prepared in 2  Tris–Bicine urea sample buffer on the gel to assign the different Aβ species. 2.1.4 MS Analysis of Aβ Species Equipment: 27-gauge syringe needle Benchtop centrifuge (1.5 mL tubes) Heating block MALDI-TOF MS instrument Microcentrifuge tubes (1.5 mL) Micropipette Micropipette tips Stainless steel MALDI sample plate Materials: Acetonitrile α-Cyano-4-hydroxycinnamic acid (CHCA) Dulbecco’s Modified Eagle Medium, high glucose, GlutaMAX™ (Thermo Scientific, Waltham, MA, USA; #61965026) (DMEM) DMSO EDTA

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FCS Hydrochloric acid (HCl) N-octylglycopyranoside PGS Sodium chloride (NaCl) Trifluoroacetic acid (TFA) Tris Antibodies: Antibody recognizing all Aβ species. For best results, mouse monoclonal antibody 4G8 against Aβ17-24 (Covance, Princeton, NJ, USA; SIG-39220) is used. Solutions: DMEM/FCS DMEM Including 10% FCS Component

Final Concentration

Stock

Amount

FCS

10%

—

50 mL

DMEM

—

—

Add to 500 mL

20 × IP-MS Buffer 200 mM Tris–HCl, pH 8.0, 2800 mM NaCl, 100 mM EDTA, 2% N-octylglycopyranoside Component

Final Concentration

Stock

Amount

Tris–HCl, pH 8

200 mM

1M

25 mL

NaCl

2800 mM

5M

70 mL

EDTA, pH 8

100 mM

0.5 M

25 mL

N-octylglycopyranoside

2%

50%

5 mL

Tris–EDTA (TE) 750 mM Tris–HCl, pH 8.0, 125 mM EDTA Component

Final Concentration

Stock

Amount

Tris–HCl, pH 8

750 mM

1M

75 mL

EDTA, pH 8

125 mM

0.5 M

25 mL

TFA stock solution: 0.6% in H2O Protocol: Preparation of Aβ sample 1. Seed HEK293/sw cells in DMEM/FCS on 10-cm dishes. Grow cells to
80% confluence.

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2. Prepare DMEM/FCS-containing GSM or DMSO vehicle control. 3. Exchange the medium with 6 mL of the GSM or vehicle solutions prepared in DMEM/FCS and incubate the cells for 16–18 h. Do not increase the DMSO concentration above 0.1%. 4. Collect overnight culture media and centrifuge (10 min, 4°C, 10,000  g) to remove cellular debris. 5. Add 1/25 volume of TE to the cleared conditioned medium. At this stage, samples can be stored at 20°C until further use. 6. Mix 1 mL of conditioned medium with 15 μL of PGS beads and 3 μL of antibody 4G8 (final concentration 3 μg/mL). 7. Shake samples at 4°C for 4 h or O/N. 8. Collect the beads by centrifugation (1 min, 4°C, 2500  g) and carefully aspirate the supernatant. 9. Wash the beads three times with 1 mL 1 IP-MS buffer and two times with 1 mL of fresh Milli-Q water (vortex, centrifuge briefly (1 min, 4°C, 2500  g), and aspirate buffer). Be careful not to lose beads during the wash steps with water. 10. After the last aspiration step, remove water completely using a 27-gauge syringe needle. At this stage, samples can be stored at 20°C until further use. Matrix preparation and MALDI-TOF measurement 1. On the day of analysis, freshly prepare a 50% acetonitrile solution containing 0.3% TFA by mixing 200 μL acetonitrile and 200 μL 0.6% TFA. 2. Add a large spatula tip of CHCA to this mixture. 3. Incubate for at least 10 min at 37°C with vigorous agitation (800 rpm) to saturate the solution with CHCA. Spin down briefly (15 s, RT, 16,000  g) to separate unresolved CHCA and use supernatant for the next step. 4. Add 10 μL of saturated CHCA solution to the immunoprecipitated samples. 5. Vortex samples briefly and centrifuge (15 s, RT, 16,000  g). 6. Spot 0.4 μL of the supernatant on the sample plate and let it dry. 7. Repeat spotting two times. 8. Analyze samples with a MALDI-TOF MS instrument in linear mode with appropriate laser intensity. Analysis: MS data can be visualized by an appropriate program (e.g., DataExplorer® Software, Applied Biosystems). The molecular masses (Da) of the peaks of

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the Aβ species calculated using PeptideMass (http://web.expasy.org/ peptide_mass/) are: Aβ37: 4074.5; Aβ38: 4131.6; Aβ39: 4230.7; Aβ40: 4329.9; Aβ41: 4443.0; Aβ42: 4514.1; Aβ43: 4615.2. To compare different measurements, each spectrum is normalized by setting the maximum of the highest Aβ peak, typically that of Aβ40, as a 100% reference point.

2.2 Target Identification of GSMs Using Photoaffinity Labeling The target sites of GSMs have been unambiguously identified by photoaffinity labeling. To this end, photoactivatable benzophenone or azido groups have been inserted into various acidic and nonacidic GSMs to allow cross-linking to their targets. All acidic and nonacidic bridged aromatic compounds of the second generation analyzed to date bind to the PS1/2 NTF (Crump et al., 2011; Ebke et al., 2011; Jumpertz et al., 2012; Ohki et al., 2011; Pozdnyakov et al., 2013; Takeo et al., 2014). In the following protocol, we describe how the molecular targets of GSMs are identified using RO-57-BpB as example (Fig. 3). 2.2.1 Photoaffinity Labeling of γ-Secretase by GSMs Equipment: 12-well plates Centrifuge tubes (50 mL) Cooling centrifuge (50 mL tubes) Heating block Microcentrifuge tubes (1.5 mL) Micropipettes Micropipette tips

Fig. 3 Structure of the photocross-linkable GSM RO-57-BpB. This derivative of the GSM RO-57 contains a photoactivatable benzophenone group and a biotin tag used to pulldown cross-linked target(s).

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Polyacrylamide gel electrophoresis and blotting equipment Potter-Elvehjem homogenizer Refrigerated benchtop centrifuge (1.5 mL tubes) Shaker (4°C and RT) Ultracentrifuge (1.5 and/or 30 mL tubes) Ultracentrifuge tubes (1.5 and/or 30 mL) UV lamp (365 nm, 8 W) Materials: ε-Amino-n-caproic acid Biotin Bis–Tris I-Block™ (Tropix®, Applied Biosystems (Thermo Scientific) , Waltham, MA, USA; T2015) Citric acid Disodium hydrogen phosphate (Na2HPO4) Dithiothreitol (DTT) EDTA Glycerol Hydrochloric acid (HCl) NP-40 Potassium chloride (KCl) Potassium dihydrogen phosphate (KH2PO4) Protease inhibitor (PI) cocktail (Complete, Roche Diagnostics, Mannheim, Germany; 04693116001) Sodium citrate (NaCitrate) Sodium dodecylsulfate (SDS) Sodium chloride (NaCl) Streptavidin-Sepharose beads 10–20% Tricine gels (Invitrogen, (Thermo Scientific), Waltham, MA, USA; EC66252BOX) Tris Tween 20 Urea Antibodies: Antibodies raised against the subunits of γ-secretase. We use the following antibodies for detection: Nicastrin: N1660 (Sigma), PS1 NTF: PS1N (Capell et al., 1997)/SIG-39194 (Covance, Princeton, NJ, USA), PS1 CTF: 5E12 (Kretner et al., 2016), PS2 NTF: 2972 (Tomita et al., 1997), PS2 CTF: BI.HF5c (Steiner et al., 1999), APH-1aS/L: 4319 (Ebke et al., 2011), and PEN-2: 1638 (Steiner et al., 2002).

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Solutions: Hypotonic Buffer 15 mM NaCitrate/Citric Acid, pH 6.4, 1 mM EDTA, PI Component

Final Concentration

Stock

Amount

NaCitrate, pH 6.4

15 mM

1.5 M

100 μL

EDTA, pH 8

1 mM

0.5 M

20 μL

PI

1

25 

400 μL

H2 O

—

—

Add to 10 mL

Adjust pH to 6.4 with citric acid. Add PI always freshly before the experiment 3 × Bis–Tris Buffer 200 mM ε-Amino-n-caproic Acid, 150 mM Bis–Tris–HCl, pH 7 Component

Final Concentration

Stock

Amount

ε-Amino-n-caproic acid

200 mM

—

13.1 g

Bis–Tris

150 mM

—

15.7 g

H2O

—

—

Add to 500 mL

Adjust pH to 7.0 with HCl Resuspension Buffer 66 mM ε-Amino-n-caproic Acid, 50 mM Bis–Tris–HCl, pH 7, 1 mM EDTA, 5% Glycerol, 10 mM DTT, PI Component

Final Concentration

Stock

Amount

Bis–Tris buffer

1

3

16.67 mL

EDTA, pH 8

1 mM

0.5 M

100 μL

DTT

10 mM

1M

500 μL

PI

1

25 

2 mL

H2 O

—

—

Add to 50 mL

Solubilization Buffer 50 mM Tris–HCl, pH 7.6, 150 mM NaCl, 2 mM EDTA, 0.2% NP-40, 1% SDS Component

Final Concentration

Stock

Amount

Tris–HCl, pH 7.6

50 mM

1M

25 mL

NaCl

150 mM

5M

15 mL

EDTA, pH 8

2 mM

0.5 M

2 mL

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NP-40

0.2%

—

1 mL

SDS

1%

20%

25 mL

H2O

—

—

Add to 500 mL

10 × PBS-T 0.5% Tween 20 in 10 × PBS, pH 7.4 Component

Final Concentration

Stock

Amount

NaCl

1370 mM

—

80 g

KCl

27 mM

—

2g

Na2HPO4

100 mM

—

14.2 g

KH2PO4

18 mM

—

2.5 g

Tween 20

0.5%

—

5 mL

H2O

—

—

Add to 1 L

Adjust pH to 7.4 before adding H2O to 1 L 1% I-Block 1% I-Block™ in PBS-T, pH 7.4 Component

Final Concentration

Stock

Amount

PBS-T, pH 7.4

1

10 

2.5 mL

I-Block™

1%

—

0.25 g

H2O

—

—

Add to 25 mL

1.7 × Urea Sample Buffer 200 mM Tris–HCl, pH 8.8, 3.33% SDS, 1.67% DTT, 20% Glycerol, 10 M Urea, 4 mM Biotin Component

Final Concentration

Stock

Amount

Tris–HCl, pH 8.8

200 mM

1M

5 mL

SDS

3.33%

20%

4.16 mL

DTT

1.67%

—

0.42 g

Glycerol

20%

100%

5 mL

Urea

10 M

—

15 g

H2O

—

—

Add to 25 mL

Add biotin to a final concentration of 4 mM from a 120 mM stock solution in DMSO directly before use

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Glycerol stock solution: 65% glycerol in H2O DTT stock solution: 1 M DTT in H2O Protocol: 1. Resuspend cells in hypotonic buffer (850 μL/10-cm dish). 2. Adjust solution to OD600 ¼ 2 using hypotonic buffer, freeze in liquid N2, and thaw on ice. 3. Disrupt cells using a Potter-Elvehjem homogenizer (10 ) on ice. 4. Centrifuge (30 min, 4°C, 2500  g) and collect the supernatant. 5. Add glycerol to a final concentration of 5% (from 65% stock solution) and DTT to a final concentration of 10 mM to the supernatant. Centrifuge (60 min, 4°C, 100,000  g) and discard the supernatant. 6. Resuspend membrane pellet in resuspension buffer (50 μL/10-cm dish). Aliquots (50–250 μL) can be flash frozen and stored at 20°C until further use. 7. For the cross-linking experiment, use 50 μL aliquot (i.e., one 10-cm dish equivalent) of the membranes per condition. Dilute membrane aliquots 1:10 with cross-linking buffer. Per cross-link reaction, take 500 μL of the diluted membranes as sample. Keep excess sample (
50 μL) for the preparation of input controls (see step 16). 8. Add 1 μM cross-linkable GSM to the cross-linking sample. Prepare control samples (see “Tip” below) in parallel. 9. Shake the samples for 60 min at 4°C in the dark (wrap sample tubes in aluminum foil). 10. Transfer the samples to a 12-well plate. For GSMs containing a photoactivatable benzophenone group, irradiate with UV lamp for 30 min at 365 nm on ice (distance between sample and UV lamp approximately 1 cm). 11. Centrifuge (30 min, 4°C, 17,000  g) to pellet down the membranes. Discard the supernatant to remove unbound cross-linkable GSM. At this step, the pellet can be flash frozen and stored at 20°C until further use. 12. Resuspend each membrane pellet in 500 μL solubilization buffer (i.e., same sample volume as in step 7) to disrupt the γ-secretase complex. 13. Prepare Streptavidin-Sepharose beads a. Suspend the Streptavidin-Sepharose beads to obtain a slurry. Take the required amount of beads for all samples (15 μL/sample) and add 1 mL 1% I-Block. Shake the mixture for 15 min at RT. b. Centrifuge (1 min, 4°C, 2500  g) and aspirate the buffer carefully avoiding the beads. c. To wash the beads, add 1 mL solubilization buffer, vortex, centrifuge (1 min, 4°C, 2500  g), and aspirate the buffer again. Wash a total of three times.

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d. Resuspend the beads in solubilization buffer (take approximately the volume of the beads) to receive a 50% bead slurry. 14. Add 15 μL beads per sample and shake for 60 min at RT. 15. Wash three times with solubilization buffer. 16. Resuspend beads in 1.7  urea sample buffer. Prepare input controls (e.g., 1, 0.5, and 0.2% of 500 μL reaction sample) using the remaining sample from step 7 and 1.7  urea sample buffer. Incubate samples and input controls for 5 min at 65°C. 17. To analyze GSM-binding to γ-secretase, split the samples, and load them on two 10–20% Tricine gels. Run the gels at 120 V. For subsequent analysis by immunoblotting, transfer the proteins of each gel on a PVDF membrane. After blotting, cut the membrane into strips at the appropriate molecular weight for the analysis of the different subunits. Analyze blot strips of gel 1 for NCT, PS1 NTF, and CTF and of gel 2 for PS2 NTF, PS2 CTF, and PEN-2. APH-1 can be analyzed by reprobing PS CTF blot strips. Depending on antibody quality, alternative antibody combinations may have to be used for the analysis of the blot strips. Tip: To set up a basic experiment, prepare four samples (Fig. 4): Two samples with cross-linkable GSM and UV irradiation, with one of these samples additionally containing noncross-linkable, parental GSM in excess (up to 100 ) over the cross-linkable GSM used for competition of binding to

Fig. 4 Identification of PS NTFs as specific targets of RO-57-BpB.b

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ensure signal specificity. Two additional control samples include one sample with UV irradiation but without cross-linkable GSM, and one sample with cross-linkable GSM but without UV irradiation. Per condition, material from one confluent 10-cm dish is required. Therefore, a basic experiment requires the preparation of membranes from at least four 10-cm dishes. As membrane aliquots can be stored at 20°C, it is more feasible to prepare a larger amount at one time.

ACKNOWLEDGMENTS We thank Karlheinz Baumann and colleagues at Roche for the kind gift of C-terminal specific anti-Aβ antibodies and GSMs as well as the collaboration in elucidating their molecular targets and mechanism of action. The contributions of Richard Page and Amelie Ebke in establishing the experimental procedures described in this chapter are gratefully acknowledged. The development of the methods described in this chapter and its preparation was supported by grants of the Deutsche Forschungsgemeinschaft (SFB596, FOR 2290).

REFERENCES Brendel, M., Jaworska, A., Herms, J., Trambauer, J., Rotzer, C., Gildehaus, F. J., et al. (2015). Amyloid-PET predicts inhibition of de novo plaque formation upon chronic γ-secretase modulator treatment. Molecular Psychiatry, 20, 1179–1187. Brockhaus, M., Grunberg, J., Rohrig, S., Loetscher, H., Wittenburg, N., Baumeister, R., et al. (1998). Caspase-mediated cleavage is not required for the activity of presenilins in amyloidogenesis and Notch signaling. Neuroreport, 9, 1481–1486. Bursavich, M. G., Harrison, B. A., & Blain, J. F. (2016). Gamma secretase modulators: New Alzheimer’s drugs on the horizon? Journal of Medicinal Chemistry, 59, 7389–7409. Capell, A., Saffrich, R., Olivo, J. C., Meyn, L., Walter, J., Grunberg, J., et al. (1997). Cellular expression and proteolytic processing of presenilin proteins is developmentally regulated during neuronal differentiation. Journal of Neurochemistry, 69, 2432–2440. Citron, M., Oltersdorf, T., Haass, C., McConlogue, L., Hung, A. Y., Seubert, P., et al. (1992). Mutation of the β-amyloid precursor protein in familial Alzheimer’s disease increases β-protein production. Nature, 360, 672–674. Crump, C. J., Fish, B. A., Castro, S. V., Chau, D. M., Gertsik, N., Ahn, K., et al. (2011). Piperidine acetic acid based γ-secretase modulators directly bind to Presenilin-1. ACS Chemical Neuroscience, 2, 705–710. Crump, C. J., Johnson, D. S., & Li, Y. M. (2013). Development and mechanism of γ-secretase modulators for Alzheimer’s disease. Biochemistry, 52, 3197–3216. Czech, C., Burns, M. P., Vardanian, L., Augustin, A., Jacobsen, H., Baumann, K., et al. (2007). Cholesterol independent effect of LXR agonist TO-901317 on γ-secretase. Journal of Neurochemistry, 101, 929–936. Czirr, E., Leuchtenberger, S., Dorner-Ciossek, C., Schneider, A., Jucker, M., Koo, E. H., et al. (2007). Insensitivity to Aβ42-lowering non-steroidal anti-inflammatory drugs (NSAIDs) and γ-secretase inhibitors is common among aggressive presenilin-1 mutations. Journal of Biological Chemistry, 282, 24504–24513. De Strooper, B., & Chavez Gutierrez, L. (2015). Learning by failing: Ideas and concepts to tackle γ-secretases in Alzheimer’s disease and beyond. Annual Review of Pharmacology and Toxicology, 55, 419–437.

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Qi-Takahara, Y., Morishima-Kawashima, M., Tanimura, Y., Dolios, G., Hirotani, N., Horikoshi, Y., et al. (2005). Longer forms of amyloid β protein: Implications for the mechanism of intramembrane cleavage by γ-secretase. Journal of Neuroscience, 25, 436–445. Selkoe, D. J., & Hardy, J. (2016). The amyloid hypothesis of Alzheimer’s disease at 25 years. EMBO Molecular Medicine, 8, 595–608. Shirotani, K., Tomioka, M., Kremmer, E., Haass, C., & Steiner, H. (2007). Pathological activity of familial Alzheimer’s disease-associated mutant presenilin can be executed by six different γ-secretase complexes. Neurobiology of Disease, 27, 102–107. Steiner, H., Romig, H., Grim, M. G., Philipp, U., Pesold, B., Citron, M., et al. (1999). The biological and pathological function of the presenilin-1 Δexon 9 mutation is independent of its defect to undergo proteolytic processing. Journal of Biological Chemistry, 274, 7615–7618. Steiner, H., Winkler, E., Edbauer, D., Prokop, S., Basset, G., Yamasaki, A., et al. (2002). PEN-2 is an integral component of the γ-secretase complex required for coordinated expression of presenilin and nicastrin. Journal of Biological Chemistry, 277, 39062–39065. Struhl, G., & Adachi, A. (2000). Requirements for presenilin-dependent cleavage of Notch and other transmembrane proteins. Molecular Cell, 6, 625–636. Sun, L., Li, X., & Shi, Y. (2016). Structural biology of intramembrane proteases: Mechanistic insights from rhomboid and S2P to γ-secretase. Current Opinion in Structural Biology, 37, 97–107. Szaruga, M., Veugelen, S., Benurwar, M., Lismont, S., Sepulveda-Falla, D., Lleo, A., et al. (2015). Qualitative changes in human γ-secretase underlie familial Alzheimer’s disease. Journal of Experimental Medicine, 212, 2003–2013. Takami, M., Nagashima, Y., Sano, Y., Ishihara, S., Morishima-Kawashima, M., Funamoto, S., et al. (2009). γ-Secretase: Successive tripeptide and tetrapeptide release from the transmembrane domain of β-carboxyl terminal fragment. Journal of Neuroscience, 29, 13042–13052. Takeo, K., Tanimura, S., Shinoda, T., Osawa, S., Zahariev, I. K., Takegami, N., et al. (2014). Allosteric regulation of γ-secretase activity by a phenylimidazole-type γ-secretase modulator. Proceedings of the National Academy of Sciences of the United States of America, 111, 10544–10549. Tomita, T., Maruyama, K., Saido, T. C., Kume, H., Shinozaki, K., Tokuhiro, S., et al. (1997). The presenilin 2 mutation (N141I) linked to familial Alzheimer disease (Volga German families) increases the secretion of amyloid β protein ending at the 42nd (or 43rd) residue. Proceedings of the National Academy of Sciences of the United States of America, 94, 2025–2030. Weggen, S., & Beher, D. (2012). Molecular consequences of amyloid precursor protein and presenilin mutations causing autosomal-dominant Alzheimer’s disease. Alzheimer’s Research & Therapy, 4, 9. Weggen, S., Eriksen, J. L., Das, P., Sagi, S. A., Wang, R., Pietrzik, C. U., et al. (2001). A subset of NSAIDs lower amyloidogenic Aβ42 independently of cyclooxygenase activity. Nature, 414, 212–216. Weggen, S., Eriksen, J. L., Sagi, S. A., Pietrzik, C. U., Golde, T. E., & Koo, E. H. (2003). Aβ42-lowering nonsteroidal anti-inflammatory drugs preserve intramembrane cleavage of the amyloid precursor protein (APP) and ErbB-4 receptor and signaling through the APP intracellular domain. Journal of Biological Chemistry, 278, 30748–30754. Winkler, E., Hobson, S., Fukumori, A., Dumpelfeld, B., Luebbers, T., Baumann, K., et al. (2009). Purification, pharmacological modulation, and biochemical characterization of interactors of endogenous human γ-secretase. Biochemistry, 48, 1183–1197. Yamasaki, A., Eimer, S., Okochi, M., Smialowska, A., Kaether, C., Baumeister, R., et al. (2006). The GxGD motif of presenilin contributes to catalytic function and substrate identification of γ-secretase. Journal of Neuroscience, 26, 3821–3828.

CHAPTER SEVEN

Probing the Structure and Function Relationships of Presenilin by Substituted-Cysteine Accessibility Method T. Tomita1 Laboratory of Neuropathology and Neuroscience, Graduate School of Pharmaceutical Sciences, The University of Tokyo, Tokyo, Japan 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 1.1 Alzheimer Disease and γ-Secretase 1.2 Molecular Basis of γ-Secretase 1.3 Mechanism of Intramembrane Cleavage by γ-Secretase 2. Principle of Cysteine-Based Structural Analyses of Presenilin 2.1 SCAM 2.2 Evaluation of Topology and Position of Cys 2.3 Effect of Reagents That Affect the γ-Secretase Activity on the Biotinylation 2.4 Cross-Linking Experiment 2.5 Effect of Genetic Mutation on the Conformation of PS1 3. Protocol 3.1 Preparation of Single/Double Cys Mutant PS1 Expressing Cell 3.2 SCAM Using Intact Cell 3.3 SCAM Using Microsome 3.4 Cross-Linking Experiments 4. Summary and Perspectives Acknowledgments References

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Abstract Presenilin is a catalytic subunit of γ-secretase, which hydrolyzes several transmembrane proteins within the lipid bilayer, together with binding cofactors such as nicastrin, Aph-1, and Pen-2. However, the structural basis as well as molecular mechanism of this unusual proteolytic process remains unknown. We have analyzed the structure and function relationships of presenilin using the substituted-cysteine accessibility method (SCAM), which enables identification of the hydrophilic environment by the accessibility of sulfhydryl reagents to cysteine residues introduced at a desired position. In Methods in Enzymology, Volume 584 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2016.10.033

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2017 Elsevier Inc. All rights reserved.

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combination with small molecule inhibitors/modulators and cross-linking experiments, we were able to identify certain residues and regions of presenilin that contribute to its intramembrane-cleaving activity. In addition, we revealed the structural dynamics of the transmembrane domains of presenilin during the formation of the complex and its proteolytic process. The SCAM provides new insights into the relationship between the structure and activity of presenilin, and is useful for probing the protein dynamics of the membrane-embedded enzymes.

1. INTRODUCTION 1.1 Alzheimer Disease and γ-Secretase Alzheimer disease (AD) is pathologically defined by extensive neuronal loss and the appearance of two types of protein aggregates; senile plaques and neurofibrillary tangles (De Strooper & Karran, 2016; Scheltens et al., 2016). Amyloid-β protein (Aβ) is a major protein component of senile plaques, and derived from its precursor protein called amyloid precursor protein (APP) by sequential cleavages by β- and γ-secretases (Tomita, 2014) (Fig. 1). In particular, the C-terminal length of Aβ, which is determined by the γ-secretase shows heterogeneity. Major Aβ species secreted from neuronal cells is Aβ40, while Aβ37, Aβ38, Aβ39, and Aβ42 are also

Aβ

Senile plaques

β-Secretase

Extracellular

γ-Secretase Intracellular

APP

PS

Fig. 1 Schematic deposition of APP and PS. APP is sequentially cleaved by β- and γ-secretase to release Aβ. PS is a catalytic subunit of γ-secretase. Circles indicate the position of FAD-linked mutations.

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released to the conditioned medium. Several lines of evidence suggest that the longer form of Aβ is a molecular culprit of AD; Aβ42 is predominantly deposited in the brains of AD patients (Iwatsubo et al., 1994), and the most toxic and aggregation-prone Aβ species (Jarrett, Berger, & Lansbury, 1993). Moreover, several mutations in genes linked to autosomal-dominant familial AD (FAD) affect the γ-secretase-mediated cleavage to increase the Aβ42 production ratio (proportion of Aβ42 to total Aβ), or the aggregation property of Aβ. These genetic data strongly support the notion that the regulation of aggregation-prone longer Aβ production is thought to be a prime target in the development of drugs against AD (Selkoe & Hardy, 2016; Sperling, Mormino, & Johnson, 2014). However, it took about 10 years to gain a comprehensive understanding of γ-secretase, which executes endoproteolysis within the membrane (Tomita, 2014). First evidence for the molecular identity of the γ-secretase was obtained from the human genetics (Sherrington et al., 1995). FAD-associated genes PSEN1 and PSEN2 encode multipass membrane proteins called presenilin 1 (PS1) and PS2, respectively. Almost all of disease-causing mutations in PSEN genes have been found to increase the Aβ42 production ratio (Borchelt et al., 1997; Tomita et al., 1997). In contrast, knockout of presenilin genes abolished the γ-secretase cleavage of APP (De Strooper et al., 1999). Moreover, a chemical biology approach revealed that transition-state analog-type γ-secretase inhibitors (GSIs) directly target the PS proteins (Esler et al., 2000; Li et al., 2000), suggesting that PS is responsible for γ-secretase-mediated intramembrane cleavage.

1.2 Molecular Basis of γ-Secretase In general, proteases hydrolyze peptide bonds using ionized water. Thus, catalytic residues are usually located within an aqueous compartment. However, no conserved protease-related domain/motif was identified in PS. Thus, the molecular function of presenilin in the γ-secretase activity has been enigmatic. Critical idea came from the molecular biological study that the atypical aspartates of PS within hydrophobic sequence in TMD6 and TMD7 are required for the proteolytic reactions by the γ-secretase (Wolfe et al., 1999). Notably, these aspartates are contained within the YD and GxGD motifs that are conserved in type 4 prepilin peptidase proteins as well as signal peptide peptidases (Steiner et al., 2000; Weihofen, Binns, Lemberg, Ashman, & Martoglio, 2002). These findings raise the idea

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that these membrane proteins are novel intramembrane-cleaving GxGDtype aspartyl proteases (Wolfe & Selkoe, 2002). However, overexpression of PS alone in eukaryotic cells was not sufficient for the reconstitution of the γ-secretase activity. Biochemical studies demonstrated that PS was incorporated in the high molecular weight fractions. These data suggest the possibility that the other interacting proteins are also involved in the γ-secretase-mediated cleavage. Extensive genetic and biochemical analyses revealed that PS interacts with novel proteins including nicastrin (Nct) (Yu et al., 2000), anterior pharynx-defective 1 (Aph-1), and presenilin enhancer 2 (Pen-2) (Francis et al., 2002), and all these proteins are required for the γ-secretase-mediated cleavage in vitro and in vivo (Takasugi et al., 2003). Finally, although several binding partner proteins of PS have been identified, coexpression of PS with these three cofactors is sufficient for the reconstitution of γ-secretase activity (Edbauer et al., 2003; Kimberly et al., 2003; Takasugi et al., 2003). Thus, the minimal form of γ-secretase that can execute intramembrane cleavage is a membrane protein complex comprised of PS, Nct, Aph-1, and Pen-2.

1.3 Mechanism of Intramembrane Cleavage by γ-Secretase γ-Secretase-mediated cleavage sites of APP correspond to amino acid residues within its transmembrane domain (TMD). Extensive biochemical analyses revealed that γ-secretase executes an endopeptidase-like cleavage followed by carboxypeptidase-like processive/successive cleavage (Tomita, 2014). First, γ-secretase endoproteolyzes the substrate at the border between the cytosol and the membrane (called as ε-cleavage). Then the remaining N-terminal hydrophobic sequence is trimmed from the cytosolic side in a processive manner by every three to four residues (called as γ-cleavage) (Qi-Takahara et al., 2005). In the case of APP processing, the initial ε-cleavage results in generation of two longer Aβ species, Aβ48 or Aβ49 (Fig. 2). Both peptides are further processed by γ-cleavage to produce the Aβ peptides with various C-terminal lengths (Aβ48-Aβ45-Aβ42-Aβ38 or Aβ49-Aβ46-Aβ43-Aβ40, respectively). Intriguingly, some FAD-linked mutations specifically impaired the trimming of longer Aβ in vitro (Chavez-Gutierrez et al., 2012). In contrast, γ-secretase modulators (GSMs) reduce Aβ42 by accelerating the γ-cleavage process (Takeo et al., 2014). These enzymatic processes suggest the dynamic conformational regulation of PS/γ-secretase during the catalytic process. However, total understanding of the mechanistic action of PS1 still remains unclear.

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A Aβ Extracellular

Intracellular

B

37

40

43

46

49

–SNKGAIIGLMVGGVVIATVIVITLVMLKKKQ– Extracellular

38

42

45

48

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Fig. 2 γ-Secretase-mediated intramembrane cleavage of APP. (A) C-terminal stub of APP is cleaved by γ-secretase at the border of cytosolic side (gray; ε-cleavage), then successively trimmed to release Aβ (black; γ-cleavage). (B) Primary sequence of APP and the positions of ε- and γ-cleavages.

2. PRINCIPLE OF CYSTEINE-BASED STRUCTURAL ANALYSES OF PRESENILIN 2.1 SCAM Structural information of the γ-secretase provides critical information how this intramembrane protease recognizes and processes the membraneembedded substrate. However, the assembled γ-secretase complex contains more than 19 TMDs, which causes difficulties in crystallization of the fully active enzyme. In addition, membrane lipids also affect its proteolytic activity (Osenkowski, Ye, Wang, Wolfe, & Selkoe, 2008). Thus, other approaches such as cysteine scanning as well as single-particle analyses of the γ-secretase complex have been performed (Tomita & Iwatsubo, 2013). In fact, recently, three-dimensional atomic structure of an intact ˚ resolution was determined by cryohuman γ-secretase complex at 3.4 A electron-microscopy and single-particle analysis, and the horseshoe-like TMD arrangement was identified (Bai, Rajendra, Yang, Shi, & Scheres, 2015; Bai, Yan, et al., 2015; Lu et al., 2014). However, it still requires the detergent-mediated solubilization and purification, which would affect the intrinsic structure on the cellular membrane. FAD-linked mutations have been identified only in PSEN1 and PSEN2 among the γ-secretase complex genes. Thus, we focused on human PS1 for

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the structure-and-function relationship analysis. To obtain the structural information of PS1 in a membrane-embedded state, we and others investigated the water accessibility of specific amino acid residues of PS1 by the substitutedcysteine accessibility method (SCAM) (Ohki et al., 2011; Sato, Morohashi, Tomita, & Iwatsubo, 2006; Sato, Takagi, Tomita, & Iwatsubo, 2008; Takagi, Tominaga, Sato, Tomita, & Iwatsubo, 2010; Takagi-Niidome, Osawa, Tomita, & Iwatsubo, 2013; Takagi-Niidome et al., 2015; Takeo, Watanabe, Tomita, & Iwatsubo, 2012; Takeo et al., 2014; Tolia, ChavezGutierrez, & De Strooper, 2006; Tolia, Horre, & De Strooper, 2008; Tominaga, Cai, Takagi-Niidome, Iwatsubo, & Tomita, 2016). SCAM has originally been used to identify the pore-lining residues of transporters and channels defined by the covalent chemical modification by sulfhydryl reagents to cysteine residue (Cys) (Frillingos, Sahin-Toth, Wu, & Kaback, 1998; Javitch, 1998; Javitch, Shi, & Liapakis, 2002; Kaback, Sahin-Toth, & Weinglass, 2001; Karlin & Akabas, 1998; Seal, Leighton, & Amara, 1998). To systematically analyze the water accessibility of several residues other than endogenous Cys, all endogenous Cys should be replaced with other residues such as serine or alanine to generate Cys-less molecule. To ascertain that replacement of endogenous Cys affect the global structure of the target protein, the activity of the Cys-less protein should be analyzed. Then we introduced back a single Cys into the Cys-less protein at several positions (single Cys mutants). One of the important tricks of this method is that substitution with Cys appears to be very well tolerated in several membrane proteins. We assume that the sulfhydryl group of Cys is in one of four environments (Fig. 3); in

Extracellular

MTSEA-biotin Cysless

2 C

1 C

C

1

2

3

4

3

Biotinylated proteins Intracellular

4

C

Fig. 3 Principle of SCAM. Cys (indicated by “C” circles) facing extracellular hydrophilic environment (2, 3) is labeled by MTSEA-biotin, while Cys that locates within the lipid bilayer (1) and cytoplasmic side (4) is not pulled down.

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the water accessible extracellular/intracellular surface, the water accessible with limited access, the lipid accessible surface, or in the protein interior. Hydrophilic sulfhydryl-specific reagents will react with much faster with ionized sulfhydryls. In contrast, Cys at the lipid accessible surface and the protein interior would not react with the reagents. Among several sulfhydryl reagents, we mainly used the methanethiosulfonate (MTS) derivatives, which form mixed disulfide bonds with cysteine residues. N-Biotinylaminoethyl methanethiosulfonate (MTSEA-biotin) contains a biotin moiety that is linked to MTS. Reaction of MTSEA-biotin with Cys results in conjugation of the biotin to the thiol in Cys through a disulfide linkage. Thus, only the molecule that contains water accessible Cys is biotinylated and purified by avidin–biotin catch system (Hofmann & Kiso, 1976) after solubilization by SDS, a procedure that disrupts the γ-secretase complex.

2.2 Evaluation of Topology and Position of Cys As MTSEA-biotin is impermeable to the plasma membrane, only extracellular Cys would be labeled in SCAM using living intact cells (Seal et al., 1998). Thus, we are unable to discriminate whether unlabeled Cys is facing the cytosolic side or embedded in the membrane. To analyze the water accessibility of intracellular Cys, membrane microsomes should be subjected to the biotinylation by MTSEA-biotin (Fig. 4). We confirmed that our preparation contains both right-side-out and in-side-out membrane orientations (Sato et al., 2006). If the residue is detected only in the microsome preparation, the Cys resides on the cytoplasmic face of the membrane. If the residue is not detected, it can be assumed that the MTSEA-biotin is unable to react with it. Thus, using these two preparations (i.e., intact cells and microsome) in SCAM, we are able to map the topology of Cys facing the hydrophilic environment (Fig. 5). We are able to examine the accessibility of Cys further by the competition experiment using membrane-impermeable MTS derivatives; the negatively charged 2-sulfonatoethyl methanethiosulfonate (MTSES), the positively charged 2-(trimethylammonium)-ethyl methanethiosulfonate (MTSET), and the sterically bulkier derivative of MTSET, 2-(triethylammonium)ethyl methanethiosulfonate (MTS-TEAE). These reagents differ somewhat in size with MTS-TEAE > MTSET > MTSES > MTSEA (Sato et al., 2006, 2008; Takagi et al., 2010). The bulkiness and/or the charge of these MTS reagents prevent them from penetrating into narrow hydrophilic spaces due to physical and/or ionic barriers (Karlin & Akabas, 1998).

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Intact cells A

Microsome

Extracellular

MTSEA-biotin MTSEA-biotin C

C C

Intracellular B

Right-side-out

Extracellular

Inside-out

MTSEA-biotin MTSEA-biotin C C

C

Intracellular

Right-side-out

Inside-out

Fig. 4 Two sample preparations for topology mapping. (A) Cys locates at extracellular side. In this case, Cys is labeled in intact cells as well as microsome. (B) Cys faces to the intracellular side. Biotinylation of Cys is detected only in microsome.

Fig. 5 Summary of SCAM in PS1 CTF. Scheme is generated from the results of Sato et al. (2008). Primary sequence of PS1 is indicated by single-letter character representing the original amino acids. Cys labeled by MTSEA-biotin is shown by colored circle. Note that several residues around catalytic aspartates (stars) are labeled by MTSEA-biotin.

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Thus, if the labeling by MTSEA-biotin is decreased by preincubation with these MTS reagents, we will infer that Cys locates in an open hydrophilic environment (Fig. 6). In contrast, if Cys faces an aqueous but restricted environment by the location in the hydrophilic environment within the lipid bilayer, the biotinylation by MTSEA-biotin is unchanged in the presence of these MTS reagents. Using these analyses, we have reached the conclusion that TMD6 and TMD7 are exposed to a hydrophilic environment within the lipid bilayer, and the catalytic aspartates resides in the water accessible, but limited, milieu (Sato et al., 2006). Consistent with this, two catalytic residues of PS1 are located on the convex side of the TM horseshoe and accessible from the cytosolic side in the structure revealed by cryo-EM single-particle analysis (Bai, Yan, et al., 2015). However, some domains of PS1, including hydrophilic loop (HL) 1, 2, 5, and 6, were not apparent in the atomic structure. Moreover, singleparticle analyses suggested the dynamic conformational changes of the γ-secretase complex (Bai, Rajendra, et al., 2015; Elad et al., 2015; Li et al., 2014). This implicates the flexibility of these domains and the other molecules in the assembled structure of the γ-secretase. Importantly, SCAM and the competition experiment can be applied to any residues that sit in the

Fig. 6 Accessibility of labeled Cys examined by competition experiment. If Cys locates in the open hydrophilic environment, the labeling by MTSEA-biotin is decreased by preincubation of MTS reagent (e.g., MTSET). Cys facing an aqueous but restricted environment, be it due to charge or location in an aqueous pocket within the lipid bilayer, will still be available for MTSEA-biotin.

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flexible domains. In fact, we analyzed the structure of the extracellular HL1 using SCAM and found the periodic pattern of labeling efficiency, while all residues were facing the extracellular side. Together with the competition and biophysical analyses, we revealed that HL1 contains a short α-helical structure associated on the membrane, which is involved in the substrate recognition (Takagi-Niidome et al., 2015).

2.3 Effect of Reagents That Affect the γ-Secretase Activity on the Biotinylation As γ-secretase is one of prime targets in the drug development against AD, several small compounds that inhibit or modulate γ-secretase activity have been identified (De Strooper & Chavez Gutierrez, 2015). However, almost all compounds were developed in mammalian cell-based or in vitro assay using membrane fractions. Thus, the molecular mechanisms whereby these compounds affect the γ-secretase activity largely unknown. Under assumptions that the compound affects the intramembrane-cleaving activity by binding to PS and affecting the conformation of the γ-secretase, we are able to probe the binding site and/or allosteric effect of the compounds using competition experiment in SCAM. For instance, transition-state analogtype GSIs bind to the catalytic site containing aspartates (Li et al., 2000), while direct biochemical evidence showing that these inhibitors target the aspartates in TMD6 and TMD7 was lacking. We found that the accessibility of Cys residues located around the catalytic aspartates was significantly reduced by the transition-state analog-type GSI (Sato et al., 2006), suggesting the competition experiments clarify the residues involved in the inhibitor binding (Fig. 7). Intriguingly, pep15 that target to the initial substrate binding site in PS1 shows distinct competition pattern, supporting the notion that binding sites of these GSIs are different. However, we are unable to exclude the possibility that the inhibitor binding affects the allosteric changes in the conformation, thereby reducing the reactivity of Cys. Thus, the combination of chemical biology experiment that identifies the binding region of the compound is also required. We routinely utilize the photoaffinity labeling technique (Hatanaka, 2015) to reveal the molecular target of GSI/GSMs (Fuwa et al., 2006; Imamura et al., 2013; Kan et al., 2003; Morohashi et al., 2006; Ohki et al., 2011, 2014; Takeo et al., 2014). In other word, we are able to apply the competition to probe the allosteric effect of the reagents. In fact, we have shown that the reactivities of some Cys are increased by GSIs as well as GSM (Ohki et al., 2011; Takagi-Niidome et al., 2013; Takeo et al., 2014; Tominaga et al., 2016).

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Fig. 7 Summary of the results of labeling competition by GSIs in PS1 CTF. Scheme is generated from the results of Sato et al. (2008). Note that labeling competition patterns (indicated by gray circles) by L-685,458 (A) and pep15 (B) are distinct. Chemical structures of GSIs are shown below the scheme.

For instance, we detected the changes in the labeling efficiency of residues around the catalytic sites facing the cytoplasmic side by one of GSMs, ST1120. However, photoaffinity labeling revealed that ST1120 directly targets to the extracellular HL1 of PS. Thus, the changes in biotinylation would reflect the conformational changes of PS by ST1120 (Takeo et al., 2014). Collectively, we are able to speculate the binding site of reagents that affect the γ-secretase activity and/or the conformational changes of PS by competition analyses.

2.4 Cross-Linking Experiment To gain more insights into the structural characteristics of PS, we applied the systematic cross-linking experiment using MTS cross-linkers as molecular rulers. This method is originally utilized in the structural analysis of P-glycoprotein by Loo and Clarke (2001). As PS undergoes endoproteolysis to generate N-terminal fragment (NTF) and C-terminal fragment (CTF) after assembly of the active γ-secretase (Thinakaran et al., 1996), we can easily detect the polypeptide corresponds to a NTF–CTF heterodimer by the appearance/increase of the holoprotein levels (Sato et al., 2006) (Fig. 8). 1,2-Ethanediyl bismethanethiosulfonate (M2M), 1,3-propanediyl

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A

B Cross-linker

–

+

–

+

NTF + CTF

NTF + CTF

C C

NTF CTF anti-PS1 NTF

NTF

anti-PS1 CTF

CTF

Fig. 8 Schematic representation of cross-linking experiment. As active PS1 is endoproteolyzed to generate NTF and CTF (A), cross-linked product is easily detected as a holoprotein (B).

bismethanethiosulfonate (M3M), 1,4-butanediyl bismethanethiosulfonate (M4M), 1,6-hexanediyl bismethanethiosulfonate (M6M), 3,6-dioxaoctane1,8-diyl bismethanethiosulfonate (M8M), 3,6,9-trioxaundecane-1,11-diyl bismethanethiosulfonate (M11M), 3,6,9,12-tetraoxatetradecane-1,14-diyl bismethanethiosulfonate (M14M), and 3,6,9,12,15-pentaoxaheptadecane1,17-diyl bismethanethiosulfonate (M17M) are sulfhydryl-to-sulfhydryl crosslinking reagents with spacer arms of 5.2, 6.5, 7.8, 10.4, 13, 16.9, 20.8, and ˚ long, respectively (Loo & Clarke, 2001). After construction of double 24.7 A Cys mutant harboring a pair of Cys located in NTF and CTF, respectively, microsome fraction of cells expressing double Cys mutant is incubated with these crosslinkers and analyzed by immunoblotting. Copper and 1,10phenanthroline, which is a redox catalyst to oxidize free sulfhydryls and form a disulfide bond in cysteines that can collide (Klco, Lassere, & Baranski, 2003), can be also applied in this experiment (Takeo et al., 2014, 2012; Tominaga et al., 2016). Again, changes in the cross-linking by the preincubation of GSI/GSM would provide the information of the binding site of the reagents and/or the conformational changes related to the intramembrane proteolysis.

2.5 Effect of Genetic Mutation on the Conformation of PS1 FAD-linked mutations of PSEN1 are identified throughout the molecule. Almost all mutations upregulate the Aβ42 production ratio. However, precise effects of the mutations are distinct; some mutations increase Aβ42, whereas certain mutants decrease Aβ40 (Chavez-Gutierrez et al., 2012; Kumar-Singh et al., 2006). This suggests that the increased Aβ42 production ratio is general pathological effect on AD, while the molecular effects of FAD-linked mutations on PS might vary. Thus, it is very important to compare the conformational changes by mutations to understand the mechanisms

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of the intramembrane cleavage by the γ-secretase. SCAM-based assays mentioned earlier using single Cys mutant carrying FAD-linked mutation would provide important molecular information in the regulation of Aβ40/Aβ42 production. We have revealed that allosteric conformational changes of the cytosolic side of TMD4 were induced by FAD-linked mutations and affected the Aβ42-generating γ-secretase activity (Tominaga et al., 2016).

3. PROTOCOL 3.1 Preparation of Single/Double Cys Mutant PS1 Expressing Cell 3.1.1 Required Materials – Dulbecco’s modified Eagle’s medium (DMEM; WAKO, Osaka, Japan, #044-29765) supplemented with 10% (v/v) FBS (JRH Biosciences, Lenexa, KS) and 1% (v/v) penicillin and streptomycin (Invitrogen, Carlsbad, CA, # 15070063) – DKO cells (Herreman et al., 2000) – pMXs-puro (Kitamura et al., 2003) – Plat-E retrovirus packaging cells (Kitamura et al., 2003) – Puromycin (Wako, Osaka, Japan, #166-23153) – Blastcidin (Wako, Osaka, Japan, #029-18701) – FuGene 6 transfection reagent (Promega, Fitchburg, WI, #E2692) – Antibody against PS1 NTF (Sato et al., 2008; Tominaga et al., 2016) – Antibody against PS1 CTF (Tomita et al., 1997, 1999) – Human/Rat β Amyloid (40) ELISA Kit Wako (Wako, Osaka, Japan, #294-62501) – Human/Rat β Amyloid (42) ELISA Kit Wako (Wako, Osaka, Japan, #290-62601) 3.1.2 Generation and Validation of Cells Expressing Cys Mutant PS1 Construction of Cys-less, single Cys, and double Cys mutants are performed by standard molecular biology technique. As endogenous PS proteins contain Cys, we recommend to use cell lacking endogenous PS1 and PS2, such as an embryonic fibroblasts derived from Psen1/Psen2 double knockout mouse (i.e., DKO cells) (Herreman et al., 2000). As transient expression of PS causes the accumulation of the holoprotein because of inaccurate folding, we strongly recommend to generate stable cell lines for each mutant. We routinely utilize the recombinant retrovirus system based on pMXs-puro vector

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(Kitamura et al., 2003; Watanabe et al., 2005) to establish the stable cell lines. It is very important to check the expression and proteolytic activity of the mutants by Aβ generation by sandwich ELISA (or equivalent method) to ascertain that the γ-secretase containing mutant PS is properly assembled.

3.2 SCAM Using Intact Cell 3.2.1 Required Materials • All MTS reagents were dissolved in DMSO at 200 mM and stored at 80°C until use – MTSEA-biotin (Toronto Research Chemicals, Toronto, Canada, #B394750) – MTS-TEAE (Toronto Research Chemicals, Toronto, Canada, #T775800) – MTSET (Toronto Research Chemicals, Toronto, Canada, #T795900) – MTSES (Toronto Research Chemicals, Toronto, Canada, #S672000) • SDS (Nacalai tesque, Kyoto, Japan, #31606-04) • Streptavidin sepharose High Performance (GE Healthcare, Piscataway, NJ, #17511301) • L-685,458 (Peptide Institute, Osaka, Japan, #4394-v) • DAPT (Peptide Institute, Osaka, Japan, #3219-v) • Pep15 (BEX, Tokyo, Japan, custom order) (Das et al., 2003) 3.2.2 Intact Cell Biotinylation 1. Stable DKO cells are grown in 6-well plates until confluency. 2. After washing cells with ice-cold PBS, cells are incubated with 1 mM MTSEA-biotin in 450 μL of PBS for 30 min at 4°C. 3. Excess reagents are removed by washing with PBS twice. 4. Labeled cells are lysed in 1 mL of 1% SDS/PBS for 1–3 h in room temperature by shaking. 5. Solubilized supernatants are incubated with 30 μL of streptavidin sepharose (50% slurry in PBS) in 1.5 mL tube for overnight in room temperature using rotary shaker. 6. Biotinylated proteins are precipitated by centrifugation at 3000 rpm for 5 min. 7. Discard the supernatant and add 500 μL of 1% SDS/PBS. 8. After vortex, the biotinylated protein is pulled down. Repeat four times. 9. Samples are eluted by boiling in 70 μL of sample buffer containing 1% 2-mercaptoethanol at 100°C for 1 min. 10. Eluates are analyzed by immunoblotting.

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For competition analyses using MTS reagents (final concentration 2 mM) or inhibitors (final concentration 1 μM for L-685,458 and pep15, 10 μM for DAPT), living cells are preincubated with chemicals for 30 min and then subjected to biotinylation by MTSEA-biotin.

3.3 SCAM Using Microsome 3.3.1 Required Materials • All MTS reagents were dissolved in DMSO at 200 mM and stored at 80°C until use – MTSEA-biotin (Toronto Research Chemicals, Toronto, Canada, #B394750) – MTS-TEAE (Toronto Research Chemicals, Toronto, Canada, #T775800) – MTSET (Toronto Research Chemicals, Toronto, Canada, #T795900) – MTSES (Toronto Research Chemicals, Toronto, Canada, #S672000) • SDS (Nacalai tesque, Kyoto, Japan, #31606-04) • Streptavidin sepharose High Performance (GE Healthcare, Piscataway, NJ, #17511301) • PBS (8 mM Na2HPO4, 2 mM NaH2PO4, 131 mM NaCl (pH 7.4)) • Homogenization buffer (10 mM HEPES, 150 mM NaCl (pH 7.4), 10% glycerol, Complete protease inhibitor cocktail (Roche Life Science, Penzberg, Germany, #11836145001)) • Polytron homogenizer (Hitachi, Tokyo, Japan) • Ultracentrifuge (Beckman, Brea, CA) • BCA assay kit (Pierce, Waltham, MA, #23225) 3.3.2 Microsomal Preparation and Biotinylation Experiments 1. Stable DKO cells expressing cysteine mutant PS1 are grown on two 15-cm dishes per single analysis. 2. After confluency, cells are scraped into PBS and resuspended in 2 mL of homogenization buffer. 3. Cells are disrupted using a Polytron homogenizer on ice. 4. Nuclei and large cell debris are pelleted by centrifugation at 1500  g for 10 min in 4°C. 5. The postnuclear supernatants are ultracentrifuged at 20,000  g for 1 h in 4°C. 6. The resultant microsomal pellets are resuspended in 0.2 mL of PBS in a 27 gauge syringe. 7. Protein concentration of this fraction is measured by BCA assay to adjust protein concentration at 1 mg/mL.

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8. 4 μL of 50 mM MTSEA-biotin in DMSO (final concentration 1 mM) is added to 200 μL of the microsome fraction. 9. 30 min incubation at 4°C. 10. The microsomes are centrifuged at 20,000  g for 5 min in 4°C. 11. The pellet is resuspended in 200 μL PBS using a syringe. 12. The microsomes are again centrifuged at 20,000  g for 5 min in 4°C to wash out free MTSEA-biotin. 13. The resultant pellets are resuspended in 1 mL of 1% SDS/PBS for 1–3 h in room temperature by shaking. 14. Labeled proteins are incubated with the streptavidin sepharose overnight and analyzed as in the intact cell biotinylation experiment (see Section 3.2.2). For competition analyses using MTS reagents (2 mM) or inhibitors (1 μM for L-685,458 and pep15, 10 μM for DAPT), microsomes are preincubated with chemicals for 15 min in 4°C and centrifuged at 20,000  g for 5 min in 4°C. The resultant pellets are resuspended in 0.2 mL of PBS, and then subjected to biotinylation by MTSEA-biotin.

3.4 Cross-Linking Experiments 3.4.1 Required Materials • All MTS reagents were dissolved in DMSO at 200 mM and stored at 80°C until use – M2M (Toronto Research Chemicals, Toronto, Canada, #E890350) – M3M (Toronto Research Chemicals, Toronto, Canada, #P760350) – M4M (Toronto Research Chemicals, Toronto, Canada, #B690150) – M6M (Toronto Research Chemicals, Toronto, Canada, #H294250) – M8M (Toronto Research Chemicals, Toronto, Canada, #D486150) – M11M (Toronto Research Chemicals, Toronto, Canada, #U787800) – M14M (Toronto Research Chemicals, Toronto, Canada, #T306250) – M17M (Toronto Research Chemicals, Toronto, Canada, #P273750) • SDS (Nacalai tesque, Kyoto, Japan, #31606-04) • Streptavidin sepharose High Performance (GE Healthcare, Piscataway, NJ, # 17511301) • N-Ethylmaleimide (SIGMA, St. Louis, MO, #E3876) • CuSO4 (Wako, Osaka, Japan, #7758-99-8) • 1,10-Phenanthroline (SIGMA, St. Louis, MO, #131377) • EDTA (SIGMA, St. Louis, MO, #E6758) • CuPhen buffer (17 mM Tris–HCl (pH 8.0) buffer containing 1 mM CaCl2, 100 mM NaCl, and Complete protease inhibitor cocktail)

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3.4.2 Cross-Linking Experiment Using MTS Cross-Linkers 1. Microsomes are prepared as mentioned in Section 3.3.2. 2. 80 μL of resuspended microsomes (1 mg protein/mL) in PBS were incubated with MTS crosslinkers (final concentration 2 mM) for 2 h at room temperature. 3. Reaction was terminated by the further addition of 5  sample buffer containing 20 mM N-ethylmaleimide. 4. After sonication, samples were then directly subjected to immunoblot analysis. For competition analyses using inhibitors (1 μM for L-685,458 and pep15, 10 μM for DAPT), microsomes are preincubated with chemicals for 15 min in 4°C and centrifuged at 20,000  g for 5 min in 4°C. The resultant pellets are resuspended in 0.2 mL of PBS, and then subjected to biotinylation by MTSEA-biotin. 3.4.3 Cross-Linking Experiment Using Copper-Phenanthroline 1. Microsomes are prepared as mentioned in Section 3.3.2. 2. Microsomes were dissolved in CuPhen buffer (1 mg protein/mL). 3. CuSO4 and 1,10-phenanthroline (final concentration at 3 and 15 mM, respectively) are added to 80 μL of samples. 4. The mixture is incubated for 10 min at 4°C. 5. Reaction was terminated by the further addition of 5  sample buffer containing 20 mM N-ethylmaleimide and 10 mM EDTA and analyzed by immunoblot analysis.

4. SUMMARY AND PERSPECTIVES SCAM and cysteine-based structural analyses are powerful methods to study structure–function relationships and the dynamics of protein function in a variety of membrane proteins. We have adapted these approaches to γ-secretase, one of the intramembrane-cleaving enzymes that are found in all forms of life and play essential roles in biology and disease. Together with advances in structural biology, we now understand that γ-secretase forms the hydrophilic cavity containing the catalytic site within the membrane. Moreover, the conformation of presenilin is dynamically regulated during the catalytic process. Further revolutionary advances in biochemical and structural analyses would provide exciting findings regarding these novel proteases in the near future.

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ACKNOWLEDGMENTS I am grateful to Drs. Bart De Strooper (KU Leuven, Belgium), Toshio Kitamura (The University of Tokyo, Japan), Gopal Thinakaran (University of Chicago), Satoshi Yokoshima, and Tohru Fukuyama (Nagoya University, Japan) for valuable reagents used in SCAM. I would like to thank Drs. Yuichi Morohashi, Chihiro Sato, Shizuka TakagiNiidome, Aya Tominaga, Ms. Kanan Morishima, and Mr. Tetsuo Cai as SCAM group members, Dr. Takeshi Iwatsubo for longtime support of our SCAM study, and our current and previous laboratory members for helpful discussions. This work was supported in part by a Grant-in-Aid for Scientific Research (A) from the Japan Society for the Promotion of Science (JSPS) [15H02492].

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CHAPTER EIGHT

A New Method to Determine the Transmembrane Conformation of Substrates in Intramembrane Proteolysis by Deep-UV Resonance Raman Spectroscopy J.W. Cooley*,1, A. Abdine†, M. Brown*, J. Chavez†, B. Lada*, R.D. JiJi*, I. Ubarretxena-Belandia†,2 *University of Missouri, Columbia, MO, United States † Icahn School of Medicine at Mount Sinai, New York, NY, United States 2 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. In Vitro Intramembrane Proteolysis Assay 2.1 Procedure for the Purification of Substrates 2.2 Procedure for the Purification of Intramembrane Proteases 2.3 Procedure for the Expression of Isotopically Labeled Enzyme 2.4 Procedure for Intramembrane Proteolysis Assay in Detergent 2.5 Procedure for the Determination of Proteolytic Profiles 3. Intramembrane Proteolysis of the Substrate Monitored by dUVRR Spectroscopy 3.1 Procedure for dUVRR Spectral Acquisition and Analysis 4. Conclusions Acknowledgment References

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Abstract We present a new method based on deep-UV resonance Raman spectroscopy to determine the backbone conformation of intramembrane protease substrates. The classical amide vibrational modes reporting on the conformation of just the transmembrane region of the substrate can be resolved from solvent exchangeable regions outside the detergent micelle by partial deuteration of the solvent. In the presence of isotopically triple-labeled intramembrane protease, these amide modes can be accurately measured to monitor the transmembrane conformation of the substrate during intramembrane proteolysis. 1

Present address: South Bay Biomics, Manhattan Beach, CA, 90266, United States.

Methods in Enzymology, Volume 584 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2016.10.030

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2017 Elsevier Inc. All rights reserved.

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1. INTRODUCTION Intramembrane proteases (Brown et al., 2000) cleave membrane protein signaling precursors within their transmembrane domain (TMD) to release active signaling products involved in genetic regulation (Li et al., 2007), development and apoptosis in eukaryotes (Cipolat et al., 2006; Kim et al., 2006), and quorum sensing and sporulation in prokaryotes (Koide et al., 2007; Stevenson et al., 2007). Various signaling proteins are activated in this fashion, including transcriptional activators in the Notch (De Strooper et al., 1999) and ErbB-4 (Ni et al., 2001) cascades, and cholesterol regulatory proteins (Rawson et al., 1997). The β-amyloid peptides associated with Alzheimer’s disease are also the product of intramembrane proteolysis (Levy-Lahad et al., 1995; Rogaev et al., 1995; Sherrington et al., 1995; Sisodia & St George-Hyslop, 2002). Intramembrane proteases can be divided into aspartic, metallo, and serine proteases (Wolfe & Kopan, 2004). Intramembrane aspartic proteases include presenilin/γ-secretase (Wolfe et al., 1999) and signal peptide peptidase (Weihofen et al., 2002) found in humans, and the archaeal presenilin homolog MCMJR1 from Methanoculleus marisnigri JR1 discovered by us (Torres-Arancivia et al., 2010). Metalloproteases are represented by site-2-protease (Rawson et al., 1997) and the serine proteases by rhomboids (Urban, Lee, & Freeman, 2001). Crystal structures of intramembrane proteases, including MCMJR1 (Li et al., 2013), CaaX protease (Manolaridis et al., 2013; Pryor et al., 2013), site-2-protease (Feng et al., 2007), and the Escherichia coli rhomboid GlpG (Ben-Shem, Fass, & Bibi, 2007; Lemieux et al., 2007; Wang, Zhang, & Ha, 2006; Wu et al., 2006), have revealed a common architecture in which their active sites are buried within the membrane, yet the assembly of transmembrane helices keeps them sequestered from the hydrophobic environment of the lipid bilayer. In a recent tour de force, the cryo-EM structure of the human presenilin/γ-secretase complex has also revealed a similar architecture for intramembrane proteases when in complex with accessory subunits (Bai et al., 2015). The TMD of intramembrane protease substrates is presumably in an α-helical conformation (Wolfe, 2009), to avoid the large free energy cost of transferring an unsatisfied hydrogen bond donor or acceptor from an aqueous to a nonpolar environment (Engelman & Steitz, 1981). However, proteases do not usually cleave intact helices, turns/ loops, or sheets. Instead, their substrate cleavage sites are in extended conformation or flexible regions where the scissile peptide bond is readily

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accessible (Hubbard, 1998; Madala et al., 2010). Thus the question of what is the conformation of the TMD of intramembrane protease substrates becomes very relevant. We have developed a new method based on deepUV resonance Raman (dUVRR) spectroscopy to determine the backbone conformation of transmembrane substrates in the absence and presence of intramembrane protease. dUVRR spectroscopy is very sensitive to conformational changes in the ensemble secondary structure of proteins (Austin, Rodgers, & Spiro, 1993; Copeland & Spiro, 1987; JiJi et al., 2006; Jordan, Eads, & Spiro, 1995; Mikhonin et al., 2005; Oladepo et al., 2011; Roach, Simpson, & JiJi, 2012), and it offers several advantages when it comes to intramembrane proteolysis: (1) This technique can provide detailed structural information about the backbone conformation of a protein in the form of several vibrational modes surrounding the amide peptide bond (Wang et al., 1991a) (Fig. 1A): amideI (C]O stretching), II (N–H bending), III (mixture of N–H, Cα–H bending and stretch, and C–N stretching), and S (a coupled in-plane bending of N–H and Cα–H). In addition, these classical amide vibrational modes are resonantly enhanced in Raman spectra when deep-UV radiation (λex < 210 nm) is used as the excitation source.

Fig. 1 (A) Classical protein backbone amide vibrational modes. (B) Sinusoidal dependence of the amideIII3 frequency and ψ angle (Mikhonin et al., 2006). The red box denotes Ψ Ramachandran angles consistent with helical geometries (only amideIII3 signal), and the blue box with angles consistent with PPII and β-strand geometries (amideIII3 + amideS signal). The white regions correspond to forbidden ψ Ramachandran angles.

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(2) The position and intensity of the observed modes are the sum of the contributions from each peptide bond as a function of the geometrical constraints imposed by the secondary structure within which they are found (Chen & Lord, 1974; Painter & Koenig, 1976). Thus, dUVRR spectra can be readily deconvoluted to resolve the content and contribution of the different secondary α-helical, β-sheet, and disordered structural elements present (Chi et al., 1998; Shashilov & Lednev, 2008, 2010; Simpson, Balakrishnan, & Jiji, 2009; Simpson et al., 2011). Strong intensities in the amideIII1,2 submodes (1280–1300 and 1310–1330 cm–1, respectively) are characteristic of canonical α-helical content. Intensities in the amideIII3 submode (1230–1270 cm–1) coupled to a lack of intensity in the amideS mode (1390–1410 cm–1) are a clear indication of the presence of backbone populations with ψ angles correlative of helical (α-helix, π-helix, or 310-helix) elements. Finally, intensities in the amideIII3 submode coupled to an increase in amideS intensity are directly proportional to polyproline II helix (PPII) and β-strand content (Austin et al., 1993; Harada & Takeuchi, 1986; Mikhonin & Asher, 2006; Wang et al., 1991a). Notably among these submodes, the amide III3 submode has been shown to be dependent on the psi (ψ) dihedral angles of the peptide backbone (Fig. 1B) (Maegawa, Ito, & Akiyama, 2005; Shashilov & Lednev, 2010; Strisovsky, Sharpe, & Freeman, 2009). This correlation allows the determination of ψ-angles within
8 degrees by measuring the amide III3 shift and using Eq. (1) (Mikhonin et al., 2006):   cm1 1 1 ϑEXT ðT  T0 Þ III3 ¼ 1256cm  54cm sin ðψ + 26°Þ  0:11 °C

(1)

(3) Not limited by macromolecular size, and as background signal from buffer, lipid, and detergent is small and easily substractable, dUVRR spectroscopy is ideally suited to measure secondary structure of membrane proteins in detergent micelles and lipid bilayers (Brown et al., 2014; Eagleburger, Cooley, & JiJi, 2014; Halsey et al., 2011a, 2011b, 2012). In addition, dUVRR spectroscopy can resolve lipid/detergent embedded vs aqueous solvated portions of a protein (Halsey et al., 2011a). (4) No protein modifications are necessary for recording dUVRR spectra, and resolving two molecular species from one another (i.e., substrate from enzyme) is feasible using isotope labeling. Here, we present approaches and procedures for dUVRR spectroscopy measurements in the context of an in vitro intramembrane proteolysis activity

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assay in detergent micelles using purified substrate and enzyme. We show how by partial deuteration of the aqueous solvent we can resolve features of the dUVRR spectra revealing just the backbone conformation of the TMD of the substrate. In addition, we show how we can readily distinguish the vibrational amide modes of the TMD of the substrate during intramembrane proteolysis by employing triple isotopically labeled enzyme.

2. IN VITRO INTRAMEMBRANE PROTEOLYSIS ASSAY Intramembrane proteolysis is a relatively slow process, and for the application of dUVRR spectroscopy, we required a relatively efficient in vitro activity assay to ensure the conversion of the majority of substrate into product in a few hours (Fig. 2A). Reported in vitro intramembrane proteolysis assays in detergent micelles using purified substrate and enzyme have relied on chimeric substrates, in which the TMD of interest was fused to a soluble protein, and the conversion of substrate to product assessed by SDSPAGE mobility assays (Baker et al., 2007; Maegawa et al., 2007; Moin & Urban, 2012; Strisovsky et al., 2009; Urban et al., 2001; Urban & Wolfe, 2005). In an analogous manner, we designed chimeric intramembrane protease substrates containing an N-terminal maltose-binding protein (MBP), followed by a unique thrombin cleavage site motif (LVPRGS) and the TMD of interest (including 20 N-terminal juxtamembrane amino acid residues), and ending with a C-terminal His6-tag for purification. We have produced chimeric substrates containing the TMD of the Drosophila melanogaster epidermic growth factor receptor ligand rhomboid-1 substrates Gurken, Keren, and Spitz (Fig. 2B, termed MBP-Gurken-TMD, MBPKeren-TMD, and MBP-Spitz-TMD) using this general design to assay intramembrane proteolysis (Torres-Arancivia et al., 2010). For comparison, we have also produced chimeras containing not only the TMD region of Gurken, Keren, and Spitz but also their soluble cytoplasmic region (Table 1, termed MBP-Gurken-TMD/CT, MBP-Keren-TMD/CT, and MBP-Spitz-TMD/ CT). Independently of the TMD employed, these types of substrate chimeras express in E. coli cells with a high efficiency, and following metal affinity purification (Fig. 2C) a yield of 3–5 mg of protein per liter of E. coli culture can be readily achieved (Torres-Arancivia et al., 2010). Both purified MCMJR1 and GlpG rhomboid cleave these substrates in DDM micelles (Fig. 2C). In agreement with previous reports (Lemberg et al., 2005; Maegawa et al., 2005; Strisovsky et al., 2009), MBP-Gurken-TMD and MBP-Spitz-TMD are almost completely cleaved by GlpG in a period of 10 h, while 60% of

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Fig. 2 (A) Different steps in our intramembrane protease activity assay. (1) The purified and detergent-solubilized chimeric substrate composed of an N-terminal MBP, a thrombin site, TMD of interest, and a His-tag is mixed with purified intramembrane protease, which is also His-tagged and solubilized in detergent. Following incubation in 0.1% DDM at 37°C for several hours the reaction is quenched and analyzed by SDS-PAGE (as in C). (2) The MBP-containing N-terminal product is purified from the flow-through of a Ni-NTA column that bound the His-tagged C-terminal product, uncleaved substrate, and intramembrane protease. (3) The MBP containing N-terminal product is proteolyzed by thrombin to yield a short peptide (in red), flanked by the thrombin and intramembrane protease cleavage sites, for mass spectrometry analysis and cleavage site determination. (B) Cleavage profiles of GlpG and MCMJR1 against the TMD (underlined) of the substrate chimeras MBP-Gurken-TMD, MBP-Keren-TMD, and MBP-Spitz-TMD. Numbering according to the original sequence of Gurken, Keren, and Spitz. (C) Coomassie stained SDS-PAGE of purified GlpG and MCMJR1 (top panel). Activity of GlpG and MCMJR1 analyzed by SDS-PAGE and visualized by Coomassie staining against MBP-Gurken-TMD (G), MBP-Keren-TMD (K), and MBP-Spitz-TMD (S). The left panel shows the purified substrates in the absence of enzyme.

MBP-Keren-TMD is cleaved during the same period. As we previously described (Torres-Arancivia et al., 2010), MBP-Gurken-TMD is also the best substrate for MCMJR1, followed by MBP-Keren-TMD and to a less degree MBP-Spitz-TMD. Thanks to the unique thrombin cleavage site motif introduced in between the MBP region and the TMD of interest, these substrates are also

Table 1 Comparison Between Observed and Calculated Masses of Rhomboid Cleavage Products Observed Mass of Products (Da) Calculated Mass of Products (Da) Substrate

Product Sequence

MBP-Gurken-TMD

MBP-PRGS–RMA

MBP-Gurken-TMD/CT

MBP-Keren-TMD

MBP-Keren-TMD/CT

MBP-Spitz-TMD

MBP-Spitz-TMD/CT

Calc. Mass

Enzyme GlpG

PA3086

Rho-1

RHBDL-2

44,860.6

44,860

a

44,854

44,822

44,829

GS–RMA

2370.6

2371.1b

2370.5

2370.1

2369.4

MBP-PRGS–RMA

44,860.6

44,845

44,880

ND

ND

GS–RMA

2370.6

2365.8

2369.3

2367.5

2371.9

MBP-PRGS–EKA

45,306.2

45,230

45,253

45,252

45,289

GS–EKA

2816.2

2812.6

2815.2

2812.7

2815.2

MBP-PRGS–EKA

45,306.2

45,284

45,307

ND

ND

GS–EKA

2816.2

2813.2

2817.4

2811.4

2812.3

MBP-PRGS–EKA

45,385.3

45,364

45,309

45,337

45,372

GS–EKA

2895.3

2894.7

2892.8

2894.2

2894.2

MBP-PRGS–EKA

45,385.3

45,377

45,284

ND

ND

GS–EKA

2895.3

2893.7

2895.0

2894.7

2895.4

Observed mass of the N-terminal product of the reaction between rhomboid and the substrate chimera. The error in this measurement is
70 Da. Observed mass of the C-terminal product of the reaction between thrombin and the N-terminal product generated by rhomboid cleavage. The error in this measurement is
4 Da. The MBP region of the products is highlighted in bold and the thrombin cleavage motif is underlined. ND stands for non determined.

a

b

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excellent to determine the proteolytic profiles of intramembrane proteases by mass spectrometry (Torres-Arancivia et al., 2010). The uncleaved substrate, C-terminal product, and enzyme (all containing His-tags) can be bound to a metal affinity resin yielding a purified N-terminal MBPcontaining product, which can be further treated with thrombin to yield two species (Fig. 2A): one corresponding to the MBP plus a linker region cleaved at the engineered thrombin site (LVPRjGS), and the other corresponding to a short water-soluble peptide flanked by the thrombin cleavage site and the intramembrane protease cleavage site. The mass of this short peptide can be accurately measured by mass spectrometry for unambiguous determination of the proteolytic profile of a given intramembrane protease. For GlpG a single cleavage site for the MBP-Spitz-TMD (A138–S139, numbering according to the original substrate sequence), MBP-Keren-TMD (Ala122–S139), and MBP-Gurken-TMD chimeras (A245–H246) was determined. These results are in excellent agreement with the proteolytic profiles for the prokaryotic rhomboids AarA, GlpG, and YqgP determined by Strisovsky et al. (2009). The same procedure also revealed identical proteolytic profiles for the prokaryotic PA3086 rhomboid from Pseudomonas aeruginosa, eukaryotic rhomboid-1 from D. melanogaster, and RHBDL-2 from Homo sapiens (Table 1). The C-terminal region of the substrates, confirming that their TMD contains the determinants for intramembrane proteolysis, did not affect these proteolytic profiles. Notably, while rhomboids cleaved these substrates in a region corresponding to the membrane/water interface, MCMJR1 cleaved in a region corresponding to the hydrophobic core of the membrane at multiple sites in MBP-GurkenTMD (L255–L256–M257–L258) and MBP-Keren-TMD (A131–L132 and L133–F134–M135) (Torres-Arancivia et al., 2010). In summary, our assay in DDM micelles allows the measurement of substrate selectivity and proteolytic profiles of serine and aspartic intramembrane proteases. In addition, substrate conversion is relatively efficient, which favors data collection for dUVRR spectroscopy. The following is a more detailed description of the specific procedures for substrate and enzyme purification, and for performing intramembrane protease assays in detergent micelles.

2.1 Procedure for the Purification of Substrates Here, we provide an extensive and detailed protocol for the production of these chimeric substrates that is based on a more succinct procedure described before (Torres-Arancivia et al., 2010). The MBP domain was amplified from plasmid H-MBP-3C (Alexandrov, Dutta, & Pascal, 2001)

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and subcloned in between NdeI and KpnI restriction sites of IPTGinducible plasmid pET-29b, while the sequence corresponding to the 20 N-terminal juxtamembrane amino acid residues and the TMD for each substrate can be cloned in between NheI and BamHI restriction sites. Freshly transform E. coli CE43 (DE3) (OverExpress) or BL21-CodonPlus® (DE3)-RIPL (Stratagene) competent cells with the pET29b plasmid encoding the substrate of interest, and inoculate 100 mL of 2xYT medium supplemented with 50 mg/mL kanamycin in a 500-mL baffled flask. Incubate overnight at 37°C with orbital shaking at 200 rpm, and next day inoculate 1 L of enriched media (composition per liter: 10 g tryptone, 5 g yeast extract, 8.5 g Na2HPO4, 3.0 g KH2PO4, 0.5 g NaCl, 1.0 g NH4Cl, 10 mg thiamine, 6.0 g glucose) containing 34 μg/mL kanamycin in a 2-L baffled flask (Bellco Glass, Inc.) with enough overnight culture to reach 0.05 OD600. Incubate culture at 37°C with orbital shaking at 200 rpm until 0.6–0.8 OD600. Remove flask from shaker, cool it down at 4°C, add IPTG to a final concentration of 0.1 mM, and incubate at 18°C with orbital shaking at 200 rpm for 18 h. Weight the cell pellet, following harvesting by centrifugation at 6000  g for 20 min at 4°C, and resuspend it in lysis buffer (50 mM Tris–HCl at pH 7.4, 300 mM NaCl, 1 mM MgSO4, 10% glycerol, 1 mM β-mercaptoethanol, 10 μM of pepstatin (Sigma-Aldrich, USA, cat# P5318), 100 μM of leupeptin (Sigma-Aldrich, USA, cat# L2884), 0.1 μM of aprotinin (Sigma-Aldrich, USA, cat# A1153), and 100 μM PMSF (RPI, USA, cat# P20270)) at a ratio of 1 g of pellet per 5 mL of lysis buffer. Homogenize the cell suspension with a loose fit dounce homogenizer to ensure that no lumps are present during the cell lysis. Incubate the cell suspension for 30 min on ice with 100 μg each of RNase (Sigma-Aldrich, USA, cat# SRE0084), DNase (Sigma-Aldrich, USA, cat# 11284932001), and lysozyme (Sigma-Aldrich, USA, cat# L6876), prior to cell lysis using a microfluidizer (microfluidics M-110P) at 1500 psi. Alternatively, cells can be lysed in a 35-mL French pressure cell operating at 11,000 psi. Upon cell debris removal by centrifugation at 40,000  g for 20 min, cytoplasmic membranes are isolated from the supernatant by ultracentrifugation at 100,000  g for 1 h at 4°C. Resuspend membranes in solubilization buffer (50 mM Tris–HCl, pH 7.4, 300 mM NaCl, 10% glycerol) containing 10 mM imidazole (1 g of membranes per 20 mL of solubilization buffer), and solubilize for 4 h at 4°C by adding the detergent Triton X-100 (Anatrace, USA, cat# T1001, from a 20% stock) up to a final concentration of 2%. Following solubilization, centrifuge at 100,000  g for 40 min at 4°C to remove insoluble material. Provided the membrane protein of interest is stable in Triton X-100, this detergent is excellent for cell membrane

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solubilization and is also cost efficient. Mix the remaining detergent solubilizate with 0.5 mL of Ni-NTA beads (Qiagen), rock for 4 h at 4°C, and pour into a disposable purification column. After the resin settles, wash with 10-bed volumes of solubilization buffer (containing 10 mM imidazole and 0.2% Triton X-100). Immediately carry out another wash with 10-bed volumes of solubilization buffer, containing 70 mM imidazole and 0.1% n-dodecyl-β-D-maltopyranoside (DDM; Anatrace, USA, cat# D310S) detergent in exchange of Triton X-100. His-tagged substrates can be finally eluted with 5-bed volumes of solubilization buffer containing 250 mM imidazole and 0.1% DDM. The elution should be collected in 0.5 mL fractions and analyzed by SDS-PAGE. Pool the best fractions and dialyze overnight at 4°C against phosphate-buffered saline (PBS) at pH 7.4 with the addition of 10% glycerol, 0.5 mM TCEP, 0.1% NaN3, and 0.1% DDM. Following dialysis, concentrate the protein to 8 mg/mL using an Amicon ultra-4 50-kDa cutoff concentrator. Measure protein concentration using nanodrop. Finally, aliquot protein, flash-freeze in liquid nitrogen, and store at –80°C. The purified chimeras can be stored either at 4°C for over a month or at 37°C for over a week without detectable degradation. As a positive control to test if the chimeras are suitable as a substrate, we usually incubate them with thrombin (Enzyme Research Laboratories, USA), which should result in their efficient degradation into an N-terminal product that upon 4–20% SDS-PAGE analysis should yield a protein band with an apparent molecular weight of 43 kDa (corresponding to the N-terminal MBP region of the substrate), and an 7-kDa C-terminal product carrying the His6-tag corresponding to the TMD and the juxtamembrane region. Detection of the former can be achieved directly by either Coomassie blue staining or immunoblotting with anti-MBP specific antibodies (NEB, Inc.).

2.2 Procedure for the Purification of Intramembrane Proteases We use the following general protocol for production of rhomboid and MCMJR1 intramembrane proteases, which depending on the target can yield 1–3 mg of protein per liter of E. coli culture. The rhomboids GlpG and PA3086 were cloned into pET-15b plasmids for IPTG-inducible expression as N-terminal His10-tag fusion proteins and expressed in CE43 (DE3) (OverExpress) E. coli cells. Rhomboid-1 and RHBDL2 were cloned into pET-28b plasmids for inducible expression as N-terminal His10-tag fusion proteins and expressed in BL21-CodonPlus® (DE3)-RIPL E. coli

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cells. MCMJR1 was cloned into a variant of pET-28b for expression as N-terminal fusion with the small ubiquitin modifying protein (SUMO; Panavas, Sanders, & Butt, 2009), preceded by a His10-tag. MCMJR1 was expressed in E. coli strain Rosetta (DE3) pLysS (Novagen, Inc.). For expression of rhomboids grow fresh transformants in 1000 mL culture in a 2-L baffled flask at 30°C in LB broth containing either 100 μg/mL ampicillin or 34 μg/mL kanamycin until their induction (OD600 ¼ 0.4) with 0.05 mM IPTG, at which point they are incubated for another 12 h at 16°C before harvesting. For expression of MCMJR1, grow fresh transformants on 800 mL scale using a 2-L baffled flask vessel to mid-log phase at 37°C in 2xTY medium with 34 μg/mL kanamycin before lowering the temperature to 18°C and inducing protein expression with the addition of IPTG to a final concentration of 0.1 mM. Allow protein expression to continue for 18 h at 18°C. Harvest cells by centrifugation at 6000  g. In the case of rhomboids, resuspend the pelleted cells (at a ratio of 1 g of cell pellet per 5 mL of buffer) in lysis buffer (50 mM Tris–HCl, pH 7.4, 700 mM NaCl, and 10% glycerol) containing protease inhibitor cocktail (100 μM PMSF, 1 μg/mL leupeptin, 1 μg/mL pepstatin, and 1 μg/mL of aprotinin). Incubate the resuspension for 30 min on ice with 100 μg each of RNase, DNase, and lysozyme and lyse the cells with a microfluidizer (microfluidics M-110P) at 1500 psi. Remove cell debris by centrifugation at 40,000  g for 20 min and isolate cytoplasmic E. coli membranes from the supernatant by ultracentrifugation at 100,000  g for 1 h at 4°C. Resuspend membranes in lysis buffer (at 5 mg/mL total protein concentration) containing 10 mM imidazole and solubilize in 2% (w/v) Triton X-100 for 4 h at 4°C. Remove insoluble material by ultracentrifugation at 100,000  g for 40 min at 4°C, and to the supernatant add 0.5 mL of Ni-NTA beads (Qiagen), followed by rocking for 4 h at 4°C. Pour the Ni-NTA resin onto a disposable column and wash with 10-bead volumes of lysis buffer, containing 30 mM imidazole and 0.2% (w/v) Triton X-100. Wash again with 10-bead volumes of lysis buffer, containing 80 mM imidazole and 0.1% (w/v) DDM, and finally elute with 5-bead volumes of lysis buffer containing 250 mM imidazole and 0.1% (w/v) DDM. In the case of rhomboid-1 and RHBDL2, dialyze the eluate from the Ni-NTA beads against buffer (50 mM Tris–HCl, pH 7.4, 700 mM NaCl, 10% glycerol, 0.1% (w/v) DDM) and store at –80°C. In the case of rhomboids, PA3086 and GlpG concentrate the protein to 500 μL using an Amicon ultra-4 50-kDa cutoff concentrator and purify by size exclusion

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chromatography using a 16/60 HR200 Superdex column (GE Healthcare) run in 50 mM Tris–HCl, pH 7.4, 700 mM NaCl, 10% glycerol, and 0.1% (w/v) DDM. Confirm the identity of each rhomboid by in-gel chymotrypsin digestion and mass spectrometry. Variants produced by site-directed mutagenesis (QuikChange site-directed mutagenesis kit, Stratagene) can be purified as described earlier. In the case of MCMJR1 resuspend (at a ratio of 1 g of cell pellet per 5 mL of buffer) cells in lysis buffer (20 mM Na–HEPES, pH 7.5, 250 mM NaCl, 1 mM MgSO4, 1 mM β-mercaptoethanol containing 0.1 μg/mL DNAse, 4 μg/mL of E-64 (Sigma-Aldrich, USA, cat# E3132), 14 μg/mL of bestatin (Sigma-Aldrich, USA, cat# B8385), and 100 μM of PMSF). Lyse cells with a microfluidizer (microfluidics M-110P) at 1500 psi and clear cell debris by centrifugation at 40,000  g for 1 h at 4°C. Isolate cytoplasmic membranes by centrifugation at 100,000  g for 1 h at 4°C. Resuspend membranes at 5 mg/mL total protein concentration in lysis buffer and solubilized with 1% (w/v) of DDM detergent. Alternatively, the crude lysate can be solubilized directly by addition of 1% (w/v) DDM at a ratio of 10:1 (wet cell mass to detergent). Carry out solubilization at 4°C under gentle rotation for 1 h with DDM. Remove insoluble material by ultracentrifugation at 148,000  g for 1 h at 4°C and the supernatant containing the detergent-solubilized membranes was supplemented with imidazole buffered at pH 7.0 to a final concentration of 50 mM and added to preequilibrated Ni-NTA resin at a ratio of 1:50 (resin to solution, v/v). After 2 h of incubation under gentle rotation pour the affinity resin onto a disposable column and wash with 4 column volumes of buffer A (20 mM Na–HEPES, pH 7.5, 250 mM NaCl, 1 mM β-mercaptoethanol) supplemented with 50 mM imidazole and 0.1% (w/v) DDM. Elute the Histidine-tagged proteins in buffer A supplemented with 250 mM imidazole and 0.1% DDM. To remove the SUMO-tag treat the eluate overnight with ULP1 (added at 1:20 protease to substrate ratio, w/w) while dialyzing against a 20 mM Na–HEPES pH 7.0 buffer containing 250 mM NaCl, 2 mM β-mercaptoethanol, and 0.05% (w/v) DDM. After repassing the mixture through preequilibrated Ni-NTA resin, histidine-tagged ULP1 protease, SUMO-tag, and any impurities are retained, while MCMJR1 can be collected in the flow-through. To increase the purity even further, concentrate the protein to under 500 μL using an Amicon ultra-4 50-kDa cutoff concentrator before loading onto a superdex 200 HR10/30 (GE Healthcare, Inc.) size exclusion chromatography column equilibrated in buffer B containing 0.05% (w/v) DDM.

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2.3 Procedure for the Expression of Isotopically Labeled Enzyme dUVRR spectroscopy measurements require triple 2H, 15N, and 13C isotopically labeled enzyme, which can be expressed using the following general protocol. Freshly transform E. coli Bl21(DE3) competent cells with an IPTG-inducible plasmid encoding the intramembrane protease of interest, and next morning inoculate 100 mL of 2xYT medium supplemented with the corresponding antibiotic in a 500-mL baffled flask. Incubate for 12 h at 37°C with orbital shaking at 200 rpm, and in the evening inoculate 1 L of M9 medium in a 2-L baffled flask with enough 2xYT culture to reach 0.1 OD600. To prepare 1 L of triple-labeled M9 medium solubilize 6.5 g of anhydrous Na2HPO4, 3.0 g of KH2PO4, and 0.5 g of NaCl in 900 mL of deuterium oxide. Add 120 mg of MgSO4, 11 mg of CaCl2, 10 μg/mL biotin, 10 μg/mL thiamine, appropriate antibiotic, 2 g of [13C6] glucose (Sigma-Aldrich, USA, cat# 310808), and 1 g of 15NH4Cl (Sigma-Aldrich, USA, cat# 299251). Filter-sterilize the labeled M9 media and use within 24 h. Incubate culture at 37°C with orbital shaking at 200 rpm until 0.6 OD600. Remove the flask from the shaker, cool it down at 4°C, add IPTG to a final concentration of 0.2 mM, and incubate at 18°C with orbital shaking at 200 rpm for 18 h. Following harvesting, purification of each target can be achieved following the protocols described earlier.

2.4 Procedure for Intramembrane Proteolysis Assay in Detergent Mix detergent-purified chimeric intramembrane protease substrates (4 μM final concentration) with detergent-purified intramembrane protease (1.5 μM final concentration) in PBS at pH 7.4 in the case of MCMJR1, and in 50 mM Tris–HCl, pH 7.4, 700 mM NaCl, and 10% glycerol in the case of rhomboid. In both cases the final DDM concentration should be 0.1%, and the 30 μL reaction mixture should also contain 2 mM β-mercaptoethanol and a protease inhibitor cocktail (100 μM PMSF, 4 μg/mL of E-64, 1 μg/mL bestatin, and 1 μg/mL of aprotinin). The choice of detergent is critical for this assay and needs to be screened in order to identify the best detergent for activity, which might not necessarily be the same detergent used for purification of substrate and enzyme. In addition, in order to avoid a large excess of detergent micelles over protein, which would effectively dilute the substrate and enzyme concentration and affect the activity, it is best to carry out the assay at detergent concentrations at or

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below 10 the critical micelle concentration. The reaction is allowed to proceed for 10 h at 37°C in a thermomixer (Eppendorf, Germany) at 400 rpm before quenching it by the addition of 5  SDS-PAGE sample loading buffer. Proteolytic activity generates an N-terminal product ( 46 kDa) containing the MBP-tag and a C-terminal product carrying the His6-tag (4 kDa) (Fig. 2A). Detection of the former is achieved directly by either Coomassie blue staining of a 4–20% SDS-PAGE gel or immunoblotting with anti-MBP-specific (1:3000 dilution, NEB Inc.) primary antibody, followed by reaction with secondary antibody (1:10,000 dilution) and visualization using chemiluminescence (Pierce).

2.5 Procedure for the Determination of Proteolytic Profiles To maximize substrate conversion into product for the determination of the proteolytic profile, we recommend the use of a final concentration of 10 μM substrate and 2 μM enzyme (in the case of MCMJR1 use the SUMO-tagged enzyme as the His-tag is necessary for the purification of the reaction product) in a 1-mL reaction volume, and to carry out the reaction for 16 h at 37°C in a thermomixer at 400 rpm. Following incubation, add 250 μL of preequilibrated Ni-NTA resin to the reaction mixture to remove undigested chimeric substrate, C-terminal cleavage product and enzyme (all His-tagged) (Fig. 2A). For mass spectrometry analysis, concentrate the flowthrough containing the N-terminal cleavage product to 50 μL using an Amicon ultra 30-kDa cutoff concentrator and precipitate for 10 min at –20°C with 1 mL of 10% trichloroacetic acid (TCA) in acetone. Collect the precipitated protein by centrifugation at 17,000  g for 10 min at room temperature and wash the pellet with 1 mL of acetone at –20°C. Solubilize the pellet in 50 mM Tris–HCl, pH 7.4, and 8 M urea and pass through an Amicon ultra 30-kDa cutoff concentrator 3  to exchange the urea for 50 mM ammonium bicarbonate at pH 7.4. Concentrate the protein to a final volume of 40 μL and measure protein concentration (should be 1 mg/mL) with nanodrop. Take a 20-μL aliquot of the concentrated protein sample, add 5 μL of 5% TCA in acetone and analyze by MALDI-TOF mass spectrometry according to established procedures. In our case, we use a Voyager DE-STR (Applied Biosystems) mass spectrometer. Incubate a second 20-μL aliquot of the concentrated protein sample with one unit of thrombin for 4 h at 4°C to release a small peptide defined by the thrombin site at the N-terminus and the MCMJR1 cleavage site at the C-terminus (Fig. 2A). Stop the reaction by adding 5 μL of 5% TCA in acetone and

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analyze by MALDI-TOF mass spectrometry. Each mass spectrum is the average from 500 measurements and calibrated with myoglobin as an internal standard. The spectra are smoothed and analyzed using the software M-over-Z (Genomic Solutions, Inc.).

3. INTRAMEMBRANE PROTEOLYSIS OF THE SUBSTRATE MONITORED BY dUVRR SPECTROSCOPY In order to resolve the dUVRR signal arising from just the TMD of the chimeric substrates, we used deuteration. In general, deuteration of the protein backbone affects three (amideII, III, and S) out of four of the primary structurally sensitive amide modes. Specifically, the coupled amideS mode is abolished, while both the amideII and III modes derived from the deuterated backbone (termed amide II0 and III0 ) are readily visible but shifted to lower frequencies and with altered intensities. These changes allow the resolution in the dUVRR spectra of deuterated vibrational features from nondeuterated ones (Mikhonin & Asher, 2005; Wang et al., 1991a, 1991b). Overall, the TMD accounts for just 7% of the chimeric substrate sequence: the substrates contain an N-terminal MBP, followed by 20 juxtamembrane amino acid residues and a C-terminal TMD. Therefore, we argued that upon partial deuteration of the aqueous solvent, we should be able to resolve the dUVRR signal from the TMD, which is surrounded by the detergent micelle, from the rest of the substrate exposed to the deuterated solvent. At 50% D2O (Sigma-Aldrich, USA, cat# 756822), the dUVRR spectrum of MBP-Gurken-TMD in 0.1% DDM recorded over a period of 3 h (Fig. 3A, black spectrum) shows that the solvent exchangeable regions of the substrate, outside the detergent micelle, become negligible compared to nondeuterated aqueous solvent (Fig. 3A, red spectrum). This observation indicates that nearly all of the water-exposed amide protons in the backbone are readily exchanged for deuterons. Specifically, the dUVRR spectrum of partially exchanged substrate displays well-resolved amideI, II, III, and S vibrational modes. The amideI (1630–1680 cm–1) region likely contains contributions from both the deuterated (MBP + juxtamembrane region) and nondeuterated (transmembrane) backbone vibrations, while the remaining amideII (1525–1550 cm–1), amideS (1390–1410 cm–1), and amideIII (1230–1330 cm–1) regions of dUVRR spectra result predominantly from the ensemble secondary structure of the nondeuterated TMD of the substrate. Notably, the spectrum of MBP-Gurken-TMD also contains discernible amideIII1, III2, and III3

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Fig. 3 (A) D2O solvent exchange promotes resolution of the dUVRR signal corresponding to the transmembrane region of the substrate MBP-Gurken-TMD in DDM micelles. dUVRR spectrum of MBP-Gurken-TMD in 0.1% DDM in nondeuterated aqueous solvent (red spectrum), and in 50% D2O (black spectrum). (B) Triple 2H, 13C, and 15N isotopic labeling of GlpG drastically shifted the maxima to lower frequencies and decreased the intensities of the amideI and amideIII modes (1674 and 1230–1320 cm–1, respectively) arising from the enzyme. dUVRR spectrum of MBP-Gurken-TMD in 0.1% DDM, and 50% D2O (red spectrum), and triple-labeled GlpG also in 0.1% DDM and 50% D2O (orange spectrum). (C) dUVRR spectrum of MBP-Gurken-TMD in the presence of isotopically labeled GlpG in 0.1% DDM and 50% D2O as a function of incubation time (red spectrum at 6 h and black spectrum at 0 h of incubation).

submodes at 1280, 1327, and 1240 cm–1, respectively, indicating that the majority of the TMD of the substrate adopts a canonical α-helical conformation in the absence of intramembrane protease. MBP-Gurken-TMD is efficiently cleaved by GlpG rhomboid between residues A245 and H246. To determine the conformation of the transmembrane region of MBP-Gurken-TMD in the presence of GlpG, we had to first resolve the vibrational modes of the TMD of the substrate from those derived from the transmembrane (unexchanged) region of GlpG. To this end, we employed GlpG that was 2H, 13C, and 15N labeled. This triple isotopic labeling drastically decreased the contribution of the amideI and amideIII modes (1674 and 1230–1320 cm–1, respectively) to the dUVRR spectrum of GlpG in 0.1% DDM and 50% D2O. Specifically, a shift in their frequency by 30 cm–1 and a decrease in intensity (Fig. 3B, orange spectrum), relative to the spectrum of MBP-Gurken-TMD (Fig. 3B, red spectrum), indicated that we should be able to distinguish between the dUVRR signal arising from the TMD of the substrate and the enzyme. Indeed, the dUVRR spectrum of MBP-Gurken-TMD in the presence of isotopically labeled GlpG in a 2:1 substrate/enzyme ratio in 0.1% DDM and 50% D2O shows that the enzyme does not significantly contribute to

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the dUVRR signal of the amideIII vibrational mode (Fig. 3C, black spectrum). In addition, as a function of time, i.e., as a function of interaction with the enzyme, a small decrease in the intensity of the amideII band at 1546 cm– 1 , but most importantly a clear intensity increase in the entire amideIII region and in particular in the amide III1 mode at cm–1, can be observed (Fig. 3C, red spectrum). These data indicate that the TMD of the substrate transitions to structures characterized by an increase in noncanonical α-helical content, as evidenced by the intensity increase in the amide III1 mode at 1260, as a result of interaction with GlpG overtime.

3.1 Procedure for dUVRR Spectral Acquisition and Analysis Once purified, exchange substrate samples into PBS (10 mM phosphate, 500 mM NaCl, pH 7.4) containing 0.1% DDM. To all dUVRR spectroscopy samples, add sodium perchlorate to a final concentration of 50 mM, and deuterium oxide to a final concentration of 50%. We recommend a concentration of 20 μM of substrate and a ratio of 2:1 substrate:enzyme for dUVRR spectroscopy experiments. Different laboratories with access to Raman instrumentation might have particular setups. In our case, dUVRR spectra were collected on a custom-built dUVRR spectrometer of analogous design to those already reported in the literature (Huang, Balakrishnan, & Spiro, 2005; Lednev et al., 2005). A tunable frequency-quadrupled titanium-sapphire laser (Coherent Inc., Santa Clara, CA), pumped by the second harmonic of a Nd:YLF laser, was used as the excitation source. The laser source is capable of generating wavelengths between 195–210 and 210–225 nm through the use of two discrete optics sets. The sample was circulated through a temperature-controlled sample chamber and reservoir, maintained at 25°C, of an in-house design using a model 75211-10 gear pump (Cole Parmer, Vernon Hills, IL). A thin film of the sample was created by flowing the solution through a 19-gauge needle and between two thin nitinol wires (0.005 in diameter) (Small Parts, Inc., Miramar, FL). The sample film was directly irradiated by the incident excitation beam. An iris located after the third turning mirror just prior to the sampling flow chamber was employed to limit the power at the sample to 0.4–0.5 mW to avoid sample degradation (Wu et al., 2003). In our experience, continuous circulation of the sample in combination with low incident power is very effective at eliminating sample degradation over the extended exposures times required for dUVRR spectral acquisition. A continuous stream of nitrogen gas was used to eliminate ambient oxygen from the sample chamber.

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The Raman scattering was collected in the 135-degree backscattering geometry and dispersed using a 1.25-m spectrometer (Horiba Jobin Yvon Inc., Edison, NJ) equipped with a 3600 grove/mm grating. The spectrometer is equipped with a back illuminated, phosphor coated, liquid nitrogen cooled Symphony CCD camera (Horiba Jobin Yvon Inc., Edison, NJ) with a chip size of 2048  512. Individual 30 s spectra were collected over 2–3 h. Individual spectra were then exported in CSV format using Synergy software (Horiba Jobin Yvon Inc., Edison NJ). The spectrum of cyclohexane and the peak positions reported in Ferraro and Nakamoto (1994) were used to calibrate the collected dUVRR spectra. New calibration spectra were collected every collection session and used to calibrate all spectra collected in each session on each day. Analysis of UVRR spectra was carried out in the Matlab environment, version 7.1 (Mathworks, Natick, MA). Cosmic rays were removed from individual spectra using a cosmic ray removal program written in-house. Spectra were then averaged to increase signal to noise. Nonlinear least-squares algorithm was used to fit spectra using a series of Gaussian (amide modes) and Lorentzian (aromatic modes) bands as described previously (Simpson et al., 2009). Amide modes can be better approximated by Gaussian line shapes, perhaps due to heterogeneities in the ϕ and ψ dihedral angles about the Cα–CO (ϕ) and HN–Cα (ψ) bonds. When the amide modes of TMD helices are better approximated by a Voigt line shape due to reduced defects in the helical II° structure, we approximate the computationally intensive Voigt line shape using a mixed Gaussian/Lorentzian line shape.

4. CONCLUSIONS This communication presents a new method based on dUVRR spectroscopy to determine the transmembrane conformation of substrates in intramembrane proteolysis. Using isotope labeling strategies, it is possible to resolve the dUVRR signal arising from the TMD of the substrate in detergent micelles in the presence of intramembrane proteases. By simply substituting the TMD of interest into the chimeric substrate, this approach can be applied to a variety of substrates and intramembrane proteases. In combination with site-directed mutagenesis, this spectroscopic approach can also be used to determine the effect of specific amino acid residues on the transmembrane conformation of the substrate and on intramembrane proteolysis. In addition, dUVRR spectroscopy is well suited to measure

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protein conformation in membranes, and the intramembrane proteolysis assay in detergent micelles described here can be adapted to purified substrate and enzyme reconstituted into synthetic proteoliposomes. Finally, this approach can also be applied to measure the conformation of nascent intramembrane proteolysis products, which depending on the site of transmembrane cleavage might remain transmembrane, or escape to the aqueous environment. Thus in summary, the dUVRR spectroscopy-based approach presented here could be a valuable tool to unravel the relation between transmembrane substrate conformation and intramembrane proteolysis.

ACKNOWLEDGMENT Funding for this work was provided by Grant MCB-1244236 from the National Science Foundation.

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CHAPTER NINE

An Inducible Reconstitution System for the Real-Time Kinetic Analysis of Protease Activity and Inhibition Inside the Membrane R.P. Baker, S. Urban1 Johns Hopkins University School of Medicine, Baltimore, MD, United States 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Enzymatic Considerations for Kinetic Analysis of Proteolysis in Detergent Micelle Systems 3. A New Strategy: Quantitative Analysis in Proteoliposomes 4. Preparation of Liposomes 5. Preparation of Fluorophore-Labeled Substrate 6. Establishing an Inducible and Fluorogenic Intramembrane Protease Assay 6.1 Coreconstitution of Intramembrane Protease and Substrate 6.2 Gel Analysis of Reconstitution Efficiency and Reaction Products 6.3 Extended Progress Curve Analysis 6.4 Substrate Orientation 6.5 Further Characterization of the Experimental System 7. Real-Time Kinetic Analysis of Membrane-Immersed Proteolysis 7.1 Steady-State Kinetic Analysis of Proteolysis in Membranes 7.2 Intramembrane Protease Inhibition Assay 8. Summary Acknowledgments References

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Abstract Intramembrane proteases are an ancient and diverse group of multispanning membrane proteins that cleave transmembrane substrates inside the membrane to effect a wide range of biological processes. As proteases, a clear understanding of their function requires kinetic dissection of their catalytic mechanism, but this is difficult to achieve for membrane proteins. Kinetic measurements in detergent systems are complicated by micelle fusion/exchange, which introduces an additional kinetic step and imposes system-specific behaviors (e.g., cooperativity). Conversely, kinetic analysis in

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proteoliposomes is hindered by premature substrate cleavage during coreconstitution, and lack of methods to quantify proteolysis in membranes in real time. In this chapter, we describe a method for the real-time kinetic analysis of intramembrane proteolysis in model liposomes. Our assay is inducible, because the enzyme is held inactive by low pH during reconstitution, and fluorogenic, since fluorescence emission from the substrate is quenched near lipids but restored upon proteolytic release from the membrane. The precise measurement of initial reaction velocities continuously in real time facilitates accurate steady-state kinetic analysis of intramembrane proteolysis and its inhibition inside the membrane environment. Using real data we describe a step-by-step strategy to implement this assay for essentially any intramembrane protease.

1. INTRODUCTION Although many important reactions are catalyzed by enzymes that are tethered to the cell membrane, proteolysis occurring directly inside the cell membrane has been discovered more recently (De Strooper et al., 1999; Rawson et al., 1997; Sakai et al., 1996; Urban, Lee, & Freeman, 2001; Weihofen, Binns, Lemberg, Ashman, & Martoglio, 2002; Wolfe et al., 1999). These enzymes evolved in four distinct families that are conserved in all forms of life from bacteria to man and act as control points for a wide range of cellular processes. The site-2 protease family of metalloproteases releases membraneanchored transcription factors to control cholesterol and fatty acid biosynthesis in humans, and virulence circuits in a broad range of bacterial species (Brown, Ye, & Rawson, 2000; Makinoshima & Glickman, 2006). Several members of the aspartyl intramembrane protease family play prominent roles in the immune system, while the γ-secretase complex is expressed ubiquitously and has been implicated in Alzheimer’s disease and certain malignancies (De Strooper & Annaert, 2010; Wolfe, 2009). Rhomboid proteases are intramembrane serine proteases that activate signaling in animals, and a mitochondrial rhomboid protease regulates mitophagy with direct implications in Parkinson’s disease (Jin et al., 2010). Parasite-encoded rhomboid proteases modulate adhesion between host and parasite in a range of pathogens including the malaria parasite (Urban, 2009). Finally, the most recently discovered Rce-1 family of glutamyl intramembrane proteases processes prenylated proteins as a final step in their maturation (Manolaridis et al., 2013). A prominent target of Rce-1 in humans is the G-protein Ras, which is one of the most commonly mutated oncogenes in all cancers. The remarkable ubiquity of intramembrane proteases (Kinch, Ginalski, & Grishin, 2006; Koonin et al., 2003; Lemberg & Freeman, 2007) and their

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numerous implications in disease processes (as reviewed in De Strooper & Annaert, 2010; Urban, 2009; Urban & Dickey, 2011) has sparked an interest in deciphering their unique catalytic mechanism as a means to understand such biomedically important enzymes. A major advance toward this goal was establishing the heterologous expression and purification of intramembrane proteases in catalytically active form (Akiyama, Kanehara, & Ito, 2004; Lemberg et al., 2005; Urban & Wolfe, 2005). Appropriate detergents (usually gentle alkyl glycosides) were instrumental to the success of these studies; detergents solubilize membrane proteins from the cell membrane and thus allow separation of intramembrane proteases away from other membrane-resident proteins of a cell. In doing so, detergents convert the complex, two-dimensional system of the membrane (for which few biochemical techniques are available) into a three-dimensional environment that allows established purification, analysis, and crystallization techniques to be applied. One model intramembrane protease in particular, the rhomboid serine protease GlpG from Escherichia coli, has been the focus of intense biochemical investigation over the past decade. Early experiments were first to establish that these proteins alone catalyze intramembrane proteolysis (Urban & Wolfe, 2005). Ultimately, applying standard crystallization techniques to active GlpG in detergent systems culminated in determination of the first intramembrane protease structure (Ben-Shem, Fass, & Bibi, 2007; Lemieux, Fischer, Cherney, Bateman, & James, 2007; Wang, Zhang, & Ha, 2006; Wu et al., 2006). Ongoing studies are continuing to yield a wealth of information through structure-based functional assays and thermodynamic analyses conducted in detergent (Baker & Urban, 2012; Paslawski et al., 2015). Despite these important advances, achieving a complete understanding of this unique class of enzymes requires careful kinetic analysis to uncover their precise catalytic properties. Indeed, since intramembrane proteases did not evolve from their soluble counterparts, we should not expect them to share identical catalytic mechanisms. In this regard, detergents should be used only as a stepping-stone to analyzing intramembrane protease catalysis in their natural membrane environment. Here we describe a method for temporarily and reversibly inactivating an intramembrane protease during reconstitution with its substrate into liposomes. This inducible system prevents the premature processing of substrate during reconstitution, allowing the precise measurement of initial reaction rates. An amino-terminal fluorescent label on the substrate is quenched by the lipid environment and becomes highly fluorescent when released by proteolytic cleavage of the

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Reaction components

Protein/detergent/ liposome mixture Legend: pH 4

Lipid

Detergent

Dilution Protease

Ultracentrifugation Proteoliposomes

ON (neutral)

OFF (acidic)

Substrate ON (fluorescent)

OFF (quenched)

Discard supernatant Resuspend pellet in neutral buffer

Monitor FITC fluorescence

Fig. 1 Schematic diagram of the inducible reconstitution and fluorogenic intramembrane protease assay. Pure rhomboid protease and FITC-TatA substrate in detergent micelles are mixed with liposomes at low pH to inactivate the protease during reconstitution. Detergent is removed from the protein/lipid/detergent complexes by means of dilution followed by ultracentrifugation. Resuspension of the proteoliposome pellet in neutral reaction buffer reactivates the protease. The FITC fluorophore on the reconstituted substrate is quenched by its proximity to the membrane lipids, allowing detection of a fluorogenic signal as evidence of proteolytic release of the FITC-labeled amino-terminus.

substrate. This fluorogenic signal enables the real-time monitoring of proteolysis occurring in the membrane and facilitates steady-state kinetic analysis. Our inducible and fluorogenic assay (outlined schematically in Fig. 1) is also amenable to the study of protease inhibition kinetics as potential

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inhibitors can easily be introduced concomitant with enzyme reactivation to evaluate potency in the actual membrane environment, which is the only natural setting for these enzymes in the cell. Finally, this strategy could be applied to any intramembrane protease.

2. ENZYMATIC CONSIDERATIONS FOR KINETIC ANALYSIS OF PROTEOLYSIS IN DETERGENT MICELLE SYSTEMS Conducting kinetic analysis of intramembrane protease catalysis in detergent systems instead of in membranes affords the major advantage of applying standard techniques to these unusual enzymes. However, micellar systems like detergents introduce several unappreciated hydrodynamic features that are not true characteristics of the enzymes, and intramembrane proteases themselves have been found to display different properties in membranes. Given the importance of these considerations, we will detail each specifically in turn (and summarized in Table 1). Kinetic rate measurements performed in detergent micelle systems are confounded by several inherent limitations. First, reactions in detergents occur as a result of micelle–micelle fission/fusion events; since substrates and enzymes are contained within separate detergent micelles, catalysis relies on the rate of exchange between different micelles (Barzykin & Tachiya, 1994) and therefore may not accurately reflect the enzyme’s rate in natural membranes. Second, the true transmembrane substrate concentration in a detergent system is the concentration of protein relative to micelles, not the protein relative to buffer solution. As such, the measured KM for the same protein concentration of substrate/enzyme changes depending on the amount and/or type of detergent used in the reaction. For example, a Michaelis–Menten kinetic analysis, performed in the presence of identical protein concentrations and buffer constituents, but 0.1% vs 0.3% n-dodecyl-β-D-maltoside (DDM) detergent (not an uncommon difference between labs), results in a twofold change in KM when measured exactly in parallel (Fig. 2). Changing the type of detergent is likely to have an even greater impact. Third, detergents by their nature are inherently cooperative systems, and thus observing cooperativity in a kinetic experiment conducted in detergent should be expected and not ascribed to the enzyme without further evidence. In fact, the degree of cooperativity changes depending on detergent concentration when all other parameters are held constant (Fig. 2). Fourth, steady-state enzyme kinetics relies on achieving a plateau

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Table 1 Limitations of Detergent Micelle Systems for Kinetic Analysis Detergent Micelle System Properties Implications for Kinetic Analysis

Sequestration: enzyme and substrate are in separate detergent micelles

Micellar fusion/fission rates introduce additional and unaccounted kinetic step

Micelle dilution effect: concentration of micelles is the effective “volume” of the reaction

Kinetic parameters scale with both type and concentration of detergent, not just concentration of proteins

Cooperativity: detergents self-associate in a strongly cooperative manner

Danger of misappropriating cooperativity to enzyme:substrate interaction

Solubility: limit of solubility imposed by maximal protein:detergent ratios

Substrate solubility limit and/or distorted micelle structure may plateau reaction rates before enzyme is truly saturated

Altered dynamics: increased protein dynamics in detergent

Cleavage at ectopic sites in substrates or even nonsubstrates

Accessibility: altered accessibility of inhibitors to enzyme active site

Overestimation of inhibitor efficacies

Altered conformation: nonnative environment changes enzyme conformation/function

Catalysis may be enhanced, diminished, or even disallowed

Enzyme bias: only some intramembrane Kinetic properties of the subset of proteases are active in detergent (many enzymes active in detergent may not be require a membrane environment) generally applicable to entire class of proteases

[FITC-TatA]

[FITC-TatA]

Full length– Product– Reactions in 0.1% DDM Vmax: KM: Hill coefficient:

2045 RFU/s 180 µM 1.68

Reactions in 0.3% DDM 2143 RFU/s 359 µM 1.36

Fig. 2 Steady-state kinetic analysis in detergent. FITC-TatA substrate titrations performed under identical reaction conditions except detergent concentration was either 0.1% (left panel) or 0.3% (right panel). While the kinetic parameter, Vmax, was unaffected, KM was approximately double in the higher detergent condition. The increased concentration of detergent also resulted in decreased cooperativity, as evidenced by the lower Hill coefficient.

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of enzyme velocity with increasing substrate as evidence of enzyme saturation. However, substrate solubility limits are often reached before saturation of enzyme with substrate can be achieved. In fact, often the reaction rate plateaus that are observed (and erroneously interpreted as enzyme saturation) result from substrate aggregation or altered detergent/protein complexes as the ratio of protein to micelle is changed (because detergent concentration is usually held constant, increasing the amount of protein alters the number of substrates in each micelle). As such, perceived KM constants in detergent systems appear much lower (tighter) than the true binding constant for a substrate (Dickey, Baker, Cho, & Urban, 2013). In addition to the inherent properties that surfactants impose, intramembrane proteases behave differently in detergent vs membrane systems: GlpG, for example, generates incorrect cleavage sites in substrates, has a 10-fold higher turnover constant (kcat), and even cleaves some nonsubstrates in detergent systems (Dickey et al., 2013; Moin & Urban, 2012). This is because the membrane environment plays an active role in “tempering” enzyme and substrate dynamics, while both are radically changed in detergent systems (Moin & Urban, 2012). Inhibitor accessibility to the active site is also altered in detergent vs membrane systems. For example, the smallmolecule inhibitor JLK6 (7-amino-4-chloro-3-methoxy-isocoumarin) has a >10-fold more potent EC50 in detergent vs membrane with exactly the same substrate/enzyme (Fig. 6A). Finally, most medically relevant rhomboid proteases are inactive in detergent systems; the only eukaryotic rhomboid protease that has been characterized biochemically, Drosophila melanogaster Rhomboid-4 (Baker & Urban, 2015), absolutely requires reconstitution for activity. Ascribing enzymatic properties to rhomboid proteases as a family based on analysis of a minority that remains active in detergents introduces a serious sampling bias.

3. A NEW STRATEGY: QUANTITATIVE ANALYSIS IN PROTEOLIPOSOMES An appealing, although challenging, alternative is to study intramembrane protease kinetics in model liposomes that better approximate the native membrane environment. Biochemical analysis of membrane protein cleavage in the context of a lipid bilayer requires coreconstitution of the protease and substrate into model liposomes. The first step in this process involves mixing detergent micelles containing each of the proteins with liposomes, which leads to partial “penetration” of the liposome membranes

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by the detergents/proteins, thereby generating lipid/protein/detergentmixed complexes (Fig. 1). Removal of the detergent allows reformation of the lipid bilayer around the incorporated membrane protein(s), and these protein-containing lipid vesicles are called proteoliposomes (Fig. 1). There are several methods for achieving detergent removal, including dialysis, dilution, gel filtration, and polystyrene bead adsorption (Rigaud & Levy, 2003). The limitation of any reconstitution protocol in its application to intramembrane proteases is its time-consuming nature. Coreconstitution of an active intramembrane protease with its substrate into liposomes inevitably results in the premature processing of substrate, thereby preventing the accurate measurement of initial reaction rates. We circumvent this problem by inactivating the protease during coreconstitution with substrate into liposomes, and subsequently reactivating the protease at the onset of kinetic analysis (Fig. 1). Inactivation is achieved by lowering the pH to 4, thereby protonating the active site histidine of GlpG. We use sodium acetate buffer (pKa ¼ 4.76) because it is an effective buffer in the low pH range and has been observed to enhance inhibition by occupying the oxyanion hole in the active site of serine proteases. Importantly, the temporary low pH treatment does not perturb the overall stability or structure of the enzyme nor does it have any residual effect on protease activity (Dickey et al., 2013). While low pH (or high pH with aspartyl proteases, for example) is a general strategy applicable to any intramembrane protease, alternative methods of protease inactivation, for example, by the inclusion of reversible smallmolecule inhibitors or the exclusion of any required cation cofactors (e.g., Zn2+ for metalloproteases) during reconstitution, should also prove effective. The ability to monitor product formation directly in “real-time” streamlines the experimental process. Our strategy in designing the FITC-TatA substrate was to use fluorescence anisotropy as a readout of proteolysis since the small, fluorescein isothiocyanate (FITC)-labeled cleavage product is expected to tumble more quickly than the full-length substrate immobilized in liposomes. However, we noticed that upon reconstitution into liposomes, the fluorescence signal emitted from the FITC-labeled TatA substrate is effectively quenched by proximity to the lipids, compared to the robust fluorescence emission in detergent micelles (Fig. 3B). Cleavage of the substrate by rhomboid proteases releases the FITC-labeled amino-terminus, thereby relieving the lipid-imposed quenching to generate a signal that can be detected in real time fluorometrically (Fig. 4B).

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A

O

O

B

OH

10,000 M E S T I A T A A F G S P

N S HN N

C

FITC fluorescence (RFU)

COOH

Cleavage site

FITC-TatA in proteoliposomes + 0.2% sarkosine

8000 6000

FITC-TatA in proteoliposomes

4000 2000 0 500

550

600

650

700

Emission wavelength (nm)

Fig. 3 Fluorogenic FITC-TatA substrate. (A) A peptide comprised of residues 1–33 of TatA is labeled amino-terminally through β-alanine linkage to fluorescein isothiocyanate. The rhomboid cleavage site between two adjacent alanine residues is indicated. (B) FITC fluorescence emission is quenched when the FITC-TatA substrate is reconstituted in proteoliposomes (black line) but robust fluorescence emission is restored when the liposomes are solubilized by 0.2% (w/v) sarcosine detergent (green line).

B

WT SAHA . Reconstitution: pre post pre post GlpG –

FITC-TatA –

Reconstitution efficiency:

~90%

~94%

C 50,000

FITC Fluorescence (RFU)

A

SAHA . Enzyme: WT pH: 4 7.4 7.4 7.4 – – + prot K: –

GlpG-SAHA + proteinase K

40,000 30,000

Full length

20,000

WT-GlpG

10,000 GlpG-SAHA

Intramembrane cleavage product Juxtamembrane cleavage products

0 0

5

10

15

20

25

30

Time (min)

Fig. 4 Key parameters for inducible intramembrane protease assay development. (A) Representative samples taken pre- and postreconstitution for reactions with either wild-type GlpG or its catalytic mutant (SAHA is GlpG-S201A + H254A) were resolved electrophoretically and imaged for GlpG protease levels by Krypton infrared protein staining followed by Odyssey infrared scanning (upper panel) and for FITC-TatA using a Typhoon fluorescence scanner (lower panel). Reconstitution efficiency of 90–95% was observed for both the protease and the substrate. (B) Real-time reaction time courses show a linear increase in FITC fluorescence over time for wild-type GlpG (black line) compared to a negligible but common “drift” in fluorescence with the catalytic mutant (red line). A robust fluorescence signal is generated when proteinase K is added to the SAHA reaction (green line). (C) SDS gel electrophoresis followed by fluorescent imaging of reaction products confirms that the FITC fluorescence signal detected for wild-type GlpG in real time corresponds to an intramembrane proteolytic cleavage product (lane 2) that is not detected when the catalytic residues are mutated (lane 3) or when the wild-type enzyme is assayed at pH 4 (lane 1). Smaller reaction products corresponding to cleavage sites outside the membrane were detected in the presence of proteinase K (lane 4).

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As a starting point, we will assume that the reader wishes to study the kinetics of a new intramembrane protease that has been purified to homogeneity in active form, and that a substrate (natural or surrogate) has been identified. We will outline our protocol with “real data” using the model rhomboid protease GlpG from E. coli, which is facile to study in this context and could even be used as an instructive parallel control.

4. PREPARATION OF LIPOSOMES Unilamellar liposomes, which are spherical vesicles comprised of a single lipid bilayer (Bangham & Horne, 1964), provide an excellent model of natural membranes and can be used to study membrane proteins. Liposomes may be custom tailored to the membrane protein under investigation to best reflect its native environment, including such parameters as lipid composition, cholesterol content, and membrane curvature (vesicle diameter). For the GlpG protease that has been expressed, detergent-solubilized, and purified from bacteria (Baker & Urban, 2012), we use liposomes prepared from a natural E. coli lipid extract to study membrane-immersed proteolysis. 1. Place 100 mg of chloroform-solubilized extract of E. coli polar lipids at 25 mg/mL (100600C, Avanti Polar Lipids, Inc., Alabaster, AL) in a round-bottom 14/20 borosilicate flask. If a defined mixture of lipids is desired, chloroform solutions of the component lipids should be mixed in the appropriate ratio during this step. This is the only way to ensure proper mixing of the lipid constituents. To produce a thin, uniform lipid film, evaporate the solvent at 37°C using a rotary evaporator (Buchi, New Castle, DE) with a digital vacuum regulator set to maintain pressure at 200 Torr. It is important that solvent is removed to completion by placing the flask under high vacuum overnight at room temperature. 2. Rehydrate lipids at a concentration of 10 mg/mL in aqueous buffer consisting of 10 mM HEPES, pH 7.3, 10 mM NaCl, and 1 mM DTT by mixing at 37°C. Complete rehydration requires extensive mixing by repeated pipetting. 3. Sonicate for 2-min intervals (in a cup horn sonicator attached to a circulating water bath set at 37°C) to produce small unilamellar vesicles, as evidenced by a change in appearance from a cloudy suspension to a uniform solution. 4. Prepare lipid vesicles of more uniform size distribution by passing the aqueous lipid solution through a Nucleopore Track-Etched Polycarbonate filter (Whatman, 800281, GE Healthcare Bio-Sciences, Pittsburgh, PA) of defined pore size (typically 30, 100, or 200 nm) using an Avanti

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Mini-Extruder (Avanti Polar Lipids, Inc., 610023) 12–14 times on a digitally controlled hotplate set to 40°C.

5. PREPARATION OF FLUOROPHORE-LABELED SUBSTRATE We use long, peptidic transmembrane substrates with amino-terminal, β-alanine linked FITC, and amidated carboxy-termini that were synthesized with standard Fmoc solid-phase chemistry and purified to >90% via reversephase HPLC. We incorporate a carboxy-terminal cysteine residue for additional site-specific labeling (e.g., membrane-impermeable probe accessibility studies, biotinylation, etc.). The twin arginine transporter A (TatA) component of Providencia stuartii is the only known natural bacterial rhomboid substrate (Stevenson et al., 2007) and is efficiently processed by many rhomboid proteases, including GlpG (Dickey et al., 2013). While the method we describe here uses a peptide corresponding to amino acids 1–33 of the 99 residues that comprise TatA naturally (Fig. 3A), other peptides based on eukaryotic rhomboid substrates (e.g., Spitz and Gurken) have been synthesized and used to study rhomboid proteolysis in our lab. We generally incorporate the entire transmembrane segment of the substrate, plus at least five amino-terminal and carboxy-terminal juxtamembrane residues, into the substrate peptide. This approach should be easily adapted to accommodate other intramembrane protease substrates of interest. 1. Resuspend the lyophilized peptide to a concentration of 200 μM in 2,2,2-trifluoroethanol (Sigma-Aldrich, St. Louis, MO) and 1 mM DTT by mixing end-over-end for 1 h at room temperature. 2. Transfer the peptide to a pear-shaped 14/20 borosilicate flask and evaporate the solvent at 37°C using a rotary evaporator (as described earlier). Place the peptide under high vacuum overnight at room temperature to remove any residual solvent. 3. Resuspend the thin peptide film to a concentration of 400 μM with a buffer comprised of 20 mM HEPES, pH 7.3, 150 mM NaCl, 1 mM DTT, and 0.2% (w/v) sodium dodecanoyl sarcosine (S300S, Anatrace, Maumee, OH). As with lipids, extensive pipetting is essential to rehydrate the peptide film. Subject the resusupended peptide to indirect sonication in a temperature-controlled cup horn device at 14°C for 5 min to disrupt any possible aggregates. Temperature control is essential to ensuring that sonication does not lead to peptide denaturation/aggregation. 4. Flash freeze substrate solutions in small (usually 20 μL) aliquots on dry ice and store at 80°C.

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6. ESTABLISHING AN INDUCIBLE AND FLUOROGENIC INTRAMEMBRANE PROTEASE ASSAY Although it is likely that our inducible and real-time assay can be applied to most, if not all, active intramembrane proteases, conditions must be carefully and deliberately set up to reap the benefits of this assay system. In fact, two broad variables require careful consideration. To date, all intramembrane proteases that have been analyzed are very slow enzymes, usually taking minutes (and even hours) to cleave each substrate molecule (Bolduc, Montagna, Gu, Selkoe, & Wolfe, 2016; Dickey et al., 2013; Kamp et al., 2015). As such, even trace “invisible” amounts of contaminating cellular proteases can create severe interference when establishing a protease assay with such slow enzymes. Second, many intramembrane proteases are dependent on a specific membrane environment to recapitulate true features of their activity. As such, testing different membrane characteristics (e.g., lipid composition, fluidity, thickness, and curvature) is ultimately important. However, a good starting point is forming liposomes from natural lipid extracts from various organisms or tissues.

6.1 Coreconstitution of Intramembrane Protease and Substrate As a starting point with a new intramembrane protease, we recommend a simple experiment to evaluate reconstitution efficiency, quenching of uncleaved substrate fluorescence by the liposomes, specificity of any detected protease activity, and effect of pH on protease activity. This can be achieved with a simple set of four reconstitution reactions: the wild-type intramembrane protease assayed at pH 7.4 and 4.0, and its catalytic mutant assayed at pH 7.4 (in the absence or presence of proteinase K as a positive cleavage control). For acid proteases, elevated pH instead of pH 4 is likely to be required; pH 8.5 works well in reversibly inactivating the aspartyl intramembrane protease γ-secretase (Bolduc et al., 2016). To evaluate reconstitution efficiency, samples of the enzyme/substrate/ liposome mixture before dilution and of the resuspended proteoliposome pellet after centrifugation are compared by quantitative gel analysis (Fig. 4A). Because these are slow enzymes, it is essential that an inactive version of the target intramembrane protease (harboring a catalytic residue mutation) be purified and analyzed in parallel (Fig. 4B). Even if no other interfering enzymes prove detectable, it is conceivable that

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time-dependent protein interactions could dequench the substrate without proteolysis actually occurring. This potential “background” can also be revealed using the catalytic mutant protein analyzed in parallel. Next, analyzing the reaction at pH 4 (Fig. 4C) will evaluate whether the pH conditions are sufficient to hold the intramembrane protease inactive (which has to be analyzed by gel, because the fluorophore is not fluorescent at pH 4 even if the proteolytic product is released from liposomes). The proteinase K control (added after resuspending the substrate-containing proteoliposomes) serves as a positive control for proteolysis generating a fluorescence signal (Fig. 4B) (and gel analysis further verifies that cleavage by proteinase K is juxtamembrane relative to intramembrane for GlpG, Fig. 4C). Finally, at the conclusion of the real-time reads, a spectral scan of the inactive mutant reaction following incubation before and after adding detergent will reveal how effectively the lipids quench noncleaved substrate fluorescence (Fig. 3B). Plate reader settings are another important consideration, but it is difficult to offer meaningful guidelines because many different instrument manufacturers and models exist. Nevertheless one point warrants emphasis: we do not use automatic gain settings, but instead manually set the photomultiplier tube (PMT) gain (to between 60 and 70 on a BioTek Synergy H4 instrument) based on prior experience or preliminary trials. This helps to ensure that relative fluorescence units (RFUs) are consistent and comparable across experiments and enzymes. 1. Mix 4–5 pmol of DDM-solubilized and purified GlpG protease or its catalytically inactive mutant (S201A + H254A) with a 100-fold excess (400 pmol) of FITC-labeled peptide substrate (prepared as described earlier) in a 1 mg/mL solution of E. coli liposomes in 50 mM NaOAc, pH 4.0, and 150 mM NaCl (30 μL final reaction volume). The final detergent concentrations with our GlpG preparations are typically 0.15–0.2 mM DDM and 0.9 mM sarcosine. Take a small (e.g., 3 μL) sample of this “prereconstitution” mixture for gel analysis. 2. After incubation for 10 min at room temperature, dilute the reaction mixture 20-fold in low salt, low pH buffer (12.5 mM NaOAc, pH 4.0, and 37.5 mM NaCl), thereby diluting both detergents well below their critical micelle concentrations to facilitate coreconstitution of the protease and substrate. Incubate the diluted solution at room temperature for 10 min. 3. Separate the proteoliposomes, which are collected in the pellet, from the detergent, which is discarded with the supernatant, by ultracentrifugation

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at 600,000  g for 30 min at room temperature. Dilution followed by ultracentrifugation is the most rapid method for detergent removal, thereby minimizing the amount of time the reaction components are maintained at low pH. 4. Resuspend the proteoliposome pellets in 75 μL neutral reaction buffer (50 mM Tris, pH 7.4, 150 mM NaCl, and 1 mM DTT) or, as a control for pH inactivation of the wild-type enzyme, in 75 μL low pH buffer (50 mM NaOAc, pH 4.0, and 150 mM NaCl). Note that resuspension of proteoliposomes requires extensive but gentle pipetting (being careful to avoid generating bubbles, which interfere with the assay). Take a “postreconstitution” gel sample to allow calculation of reconstitution efficiency (see Section 6.2). After resuspension in neutral buffer, supplement one of the two reactions containing the catalytically inactive mutant with a small amount (2.5 pmol) of proteinase K as a positive control for fluorescence signal generation. 5. For each reaction, transfer a 50 μL volume into a separate well of an optical-bottom, black 384-well plate (Nunc, 242764, Thermo Fisher Scientific, Waltham, MA) that has been prewarmed and maintained at 37°C in a digital heatblock (97043-610, VWR, Radnor, PA). Seal plates with polyester microplate sealing film (PCR-SP-S, Axygen Scientific, Union City, CA) to prevent evaporation. 6. Read fluorescence each minute over a 30 min time-course at 37°C using a Synergy H4 microplate reader (Biotek, Winooski, VT) with optics configured to read from the bottom of the plate, exciting at 480  20 nm and detecting emission at 520  20 nm, and an appropriate (60–70) manual PMT gain setting.

6.2 Gel Analysis of Reconstitution Efficiency and Reaction Products Several factors affect the efficiency of protein reconstitution into the liposome membrane, including excess detergent or incomplete pelleting. As such, the reconstitution efficiency needs to be evaluated directly by gel electrophoresis, and optimized if low. An important consideration when developing any protease activity assay based on the generation of a fluorogenic signal is to verify “activity” using a direct method to detect proteolysis. Electrophoretic separation of substrate and product is an excellent way to ensure that the fluorescence signal detected truly corresponds to product formation (Fig. 4C). This is particularly relevant when analyzing a new enzyme or substrate, or in relating the

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decreased fluorescence signal in the presence of a potential inhibitor to a decrease in substrate processing. 1. At the end of a protease reaction time-course (or with timepoints analyzed in parallel, see Section 6.4), quench a sample of each reaction by the addition of an equal volume of 2  LDS sample buffer supplemented with 1/10 volume of 10  sample reducing agent (Life Technologies, Carlsbad, CA). Samples taken pre- and postreconstitution are similarly treated. 2. Heat samples at 70°C for 10 min and then load 5–10 μL on a 12% BOLT Bis–Tris gel (Life Technologies). 3. Resolve full-length FITC-TatA substrate from the smaller proteolytic cleavage product by electrophoresis at 150 V for 40 min or until the dye front is near the bottom of the gel. 4. Perform fluorescence imaging of the gel on a Typhoon FLA 9500 imager (GE Healthcare, Marlborough, MA) using the 473 nm (blue) laser to detect the FITC-labeled substrate. GlpG protease is detected by staining the gel with Krypton infrared protein stain followed by scanning in the 700 nm channel of an Odyssey infrared scanner (Li-Cor Biosciences, Lincoln, NE). 5. Quantify bands corresponding to the full-length substrate and proteolytic cleavage product using ImageQuant TL (GE Healthcare) or similar densitometry software, and GlpG levels using Image Studio software (Li-Cor Biosciences).

6.3 Extended Progress Curve Analysis Two characteristics of the reconstituted substrate must be examined in detail before a kinetic analysis can be performed: whether all of the substrate is cleavable, and whether there is a substrate orientation bias (addressed in Section 6.4). A significant consideration for any proteolytic system is whether all of the substrate is accessible to the enzyme for cleavage. This is most readily examined by extending the progress curves until all of the substrate has had a chance to be cleaved (for slow enzymes like intramembrane proteases this may require overnight incubation). A subpopulation of substrate that cannot be cleaved could result from substrate aggregation, trapping inside the liposome lumen, or highly biased intramembrane protease reconstitution orientation bias (which we have never observed). Such a subpopulation cannot interact productively with the enzyme, and it would therefore be incorrect to include all of the substrate in calculating KM.

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1. Incubate a larger intramembrane protease reconstitution reaction for extended reaction time at 37°C, and remove and quench a series of aliquots (taken from several hours to overnight) by addition to an equal volume of 2  LDS reducing sample buffer. Heat and electrophoretically resolve the terminated reactions (as described earlier). Observing cleavage proceeding to completion may require using more enzyme in the reaction than is appropriate for kinetic assays. 2. Perform fluorescence imaging using a Typhoon fluorescence imager (as described) and quantify cleavage products for the time-course of extended reactions. Reactions in which the full-length substrate is essentially undetectable and only a low molecular weight band corresponding to the rhomboid cleavage product is visible provide evidence of complete turnover of the FITC-TatA substrate. Conversely, a plateau of cleavage where full-length, uncut substrate remains unchanged with increasing time suggests that a subpopulation of the substrate may be refractory to cleavage by the protease.

6.4 Substrate Orientation A second key consideration is substrate orientation in proteoliposomes, since nonrandom substrate orientation, although rare, is possible and could affect the actual concentration of substrate that the intramembrane protease experiences. This we assess by labeling a C-terminal cysteine residue with a membrane-impermeable IRdye-maleimide (relative to labeling in the presence of detergents that solubilize the liposomes and allow complete substrate labeling). Following gel electrophoresis, this analysis revealed that half of the FITC-TatA molecules are positioned in the liposomes with the N-terminus facing in, indicating a lack of orientation bias (Dickey et al., 2013). 1. To address whether there is substrate orientation bias in proteoliposomes, incubate reconstituted FITC-TatA resuspended in DTT-free buffer (50 mM Tris, pH 7.4, 150 mM NaCl) with a 10-fold molar excess (4000 pmol) of membrane-impermeable IRDye 800CW maleimide (Li-Cor Biosciences, 929-80020) for 2 h at RT, protected from light, in either the absence or the presence of 0.2% sarcosine detergent. 2. Quench labeling reactions by the addition of an equal volume of 2  Tricine sample buffer and 1/10 volume of NuPAGE sample reducing agent (Life Technologies), and load 5 μL samples on a 16% Tricine gel (Life Technologies, EC66955BOX) followed by electrophoresis at

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120 V for about 2 h or until the dye front has reached the bottom of the gel. 3. Detect IRDye-labeled FITC-TatA in the 800 nm channel using an Odyssey infrared imaging system and quantify with Image Studio software (Li-Cor Biosciences). 4. The amount of substrate labeled with the membrane-impermeable dye in the absence of detergent corresponds to those molecules with a dyeaccessible carboxy-terminus facing outside the proteoliposome. This amount divided by the total amount of substrate labeled (in the presence of 0.2% sarcosine) gives the fraction of substrates with outward-facing carboxy-termini, and if this fraction is close to one half, there is no orientation bias.

6.5 Further Characterization of the Experimental System While the series of experiments described earlier should establish the minimal requirements for an inducible and real-time fluorescence assay strategy, ultimately additional controls should be conducted to characterize the system in greater detail. These include verifying the time-course of product generation by electrophoresis (see Section 6.2), fluorescence generation being blocked in the presence of a known inhibitor and/or by cleavage-site substrate mutations, and mass spectrometry analysis of the cleaved fragments to verify that they correspond to the bona fide protease cleavage site. Finally, any minor effect of the pH switch on the enzyme, if any, should be evaluated for effects on protein structure, stability, and/or enzyme activity. Examples of these analyses with GlpG and experimental methods required for their execution have been published in Dickey et al. (2013).

7. REAL-TIME KINETIC ANALYSIS OF MEMBRANEIMMERSED PROTEOLYSIS 7.1 Steady-State Kinetic Analysis of Proteolysis in Membranes A final requirement before a kinetic experiment can be conducted is to determine the appropriate concentration of enzyme to use in the reactions. In general, the amount of enzyme should be titrated so that 4 h, though typically time points are taken at 2 h. The reactions are quenched with SDS-loading dye, and cleavage products are separated on a 16.5% tris-tricine gel. The fluorescein-tagged substrate and cleavage product are visualized with a fluorescence scanner (Typhoon or Storm imager from GE). The bands are analyzed by densitometry and quantified against a standard curve of substrate run concurrently on each gel.

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4.2 Verification of Enzyme and Substrate Proteoliposome Incorporation 1. Proper incorporation of substrate should be measured by fluorescence quenching of the fluorophore, circular dichroism (CD), selective cysteine labeling of substrate, and by quantifying the amount of substrate that pellets with the vesicles during the ultracentrifugation step. a. Fluorescence quenching: Vesicle incorporation of the fluoresceinlabeled substrate will cause quenching of the fluorophore (Bolduc et al., 2016; Dickey et al., 2013). After the ultracentrifugation of proteoliposomes from step 2 above, the pelleted proteoliposomes are resuspended in either 50 mM HEPES, pH 7.0, 150 mM NaCl, or 50 mM HEPES, pH 7.0, 150 mM NaCl containing 0.25% NP40 to break apart the proteoliposomes. The solutions are then excited at 485 nm and the emission spectrum measured from 510 to 700 nm for the fluorescein-labeled N43 substrate on a Synergy H4 plate reader (BioTek, Winooski, VT). The NP40-containing sample of dissolved proteoliposomes should have a much higher fluorescence intensity compared to the quenched, proteoliposomeincorporated substrate. b. CD: The substrate, comprised mostly of the transmembrane domain of notch, yields a CD spectrum consistent with an α-helical secondary structure. Proteoliposomes are made similar to the above protocol with the only exception being they are made in the presence of 10 mM Tris base pH 7.0 rather than the HEPES- and NaClcontaining buffer. The final concentration of N43 peptide is 0.2 mg/mL. CD spectra can be obtained on a standard spectropolarimeter (e.g., Jasco J-815), scanning from 190 to 270 nm. c. Cysteine labeling to determine substrate orientation: Given that substrate can incorporate into the proteoliposome in two possible orientations (N-terminus inside or N-terminus outside of the proteoliposome), the relative abundance of each orientation must be experimentally measured. This allows for the determination of the amount of substrate that is available for cleavage of substrate by γ-secretase. This is accomplished for N-terminal Cys-containing N41 substrate with a membrane-impermeable Cys reactive dye (800cw maleimide, Licor, Lincoln, NE, product number: 929–80020). 800cw is added to the formed proteoliposomes at a concentration of 1 μM in the presence or absence of 1 mg/mL melittin peptide (Sigma-Aldrich, product number: M2272) to permeabilize

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the proteoliposomes and allow passage of the dye through the membrane. The labeling reaction proceeds for 2 h at room temperature. Unreacted dye is quenched with 50 mM DTT for 30 min before the labeled peptide is run on a 16.5% tris-tricine gel and visualized with an Odyssey Licor scanner. In this way, the fraction of substrate labeled on the outside of the lipid bilayer can be measured. Fortuitously, the ratio happens to be approximately 50:50 for N41 substrate. This same experiment was carried out with FITC-TatA for rhomboid using detergent instead of melittin to permeabilize the membrane to measure total labeling (Dickey et al., 2013). d. Total substrate incorporation: Even after diluting the reaction mixture below the detergent CMC, not all of the substrate will be incorporated into the proteoliposome. This can be measured by residual fluorescence remaining in the supernatant after ultracentrifugation. To determine the amount of N43 substrate incorporated, a small aliquot of reaction mixture is taken before and after ultracentrifugation and quantified by densitometry after running on a 16.5% tris-tricine gel. The percent of substrate in the pellet is assumed to be incorporated into the proteoliposome. The percent of substrate incorporation into the proteoliposome should not vary significantly with the amount of substrate added. 2. The concentration of active γ-secretase enzyme in the proteoliposome after detergent dilution, ultracentrifugation, and subsequent resuspension in a buffer of neutral pH is determined by titrating active enzyme with a tight-binding γ-secretase transition-state analog inhibitor. LY411,575 (Sigma-Aldrich, product number: SML0506), a very potent γ-secretase inhibitor, is added to the resuspension buffer at varying concentrations, and the reaction is allowed to proceed for 2 h at 37°C. The reactions are then quenched and product visualized and quantified as described earlier. Unlike with N43 substrate, the orientation of γ-secretase after being incorporated into the proteoliposome does not need to be determined. Here, only the enzyme molecules that are oriented such that their active sites are exposed to the pH change will be active. The concentration of active enzyme molecules is determined by inhibitor titration.

5. CONCLUSIONS I-CLiPs represent a unique class of enzyme at the forefront of biology and medicine. A thorough understanding of the intramembrane protease

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catalytic mechanism will be required to fully understand how these proteases govern important biological processes as well as allow researchers to capitalize on the therapeutic potential of these enzymes. This can only be accomplished with robust and facile methods by which to study the activity of these enzymes in vitro. Although the mysteries of the intramembrane protease cleavage mechanism are only now beginning to be unraveled, the enzymatic assays outlined here have proven highly valuable for interrogating the function of γ-secretase and other members of the I-CLiP family.

ACKNOWLEDGMENTS This work was supported by NIH Grant P01 AG15379.

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CHAPTER THIRTEEN

Methods for Structural and Functional Analyses of Intramembrane Prenyltransferases in the UbiA Superfamily Y. Yang*, N. Ke†, S. Liu*, W. Li*,1 *Washington University School of Medicine, St. Louis, MO, United States † New England Biolabs, Ipswich, MA, United States 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction to the UbiA Superfamily 2. Biological Function and Enzymatic Activity of the Subfamilies 3. Predicting the Functional Role of Intramembrane Prenyltransferases by Sequence Clustering Analysis 3.1 Selecting Representative Proteins of Each Major Subfamily 3.2 Sequence Clustering Analysis 4. Analysis of Enzymatic Activity 4.1 Generation of a ubiA–menA– BL21(DE3) Strain 4.2 Complementation Assay of UbiA Activity With the ubiA–menA– BL21(DE3) Strain 4.3 Assay of Enzymatic Activity in Microsomes 5. Structural Studies of the UbiA Superfamily 5.1 Cloning, Expression, and Selection of Protein Constructs 5.2 Expression of the ApUbiA Protein in a Minimal Media Used for SeMet Labeling 5.3 Protein Purification 5.4 Selection of Detergents 5.5 Crystallization and Structure Determination 5.6 Structure Comparison of Intramembrane and Soluble Prenyltransferases Acknowledgments References

Methods in Enzymology, Volume 584 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2016.10.032

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2017 Elsevier Inc. All rights reserved.

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Abstract The UbiA superfamily is a group of intramembrane prenyltransferases that generate lipophilic compounds essential in biological membranes. These compounds, which include various quinones, hemes, chlorophylls, and vitamin E, participate in electron transport and function as antioxidants, as well as acting as structural lipids of microbial cell walls and membranes. Prenyltransferases producing these compounds are involved in important physiological processes and human diseases. These UbiA superfamily members differ significantly in their enzymatic activities and substrate selectivities. This chapter describes examples of methods that can be used to group these intramembrane enzymes, analyze their activity, and screen and crystallize homolog proteins for structure determination. Recent structures of two archaeal homologs are compared with structures of soluble prenyltransferases to show distinct mechanisms used by the UbiA superfamily to control enzymatic activity in membranes.

1. INTRODUCTION TO THE UbiA SUPERFAMILY The intramembrane prenyltransferases of UbiA superfamily generate the basic skeleton of a variety of lipophilic compounds. These prenyltransferases catalyze the fusion of an isoprenyl or phytyl chain to another molecule, often an aromatic compound from the aqueous phase of cells. Addition of the aliphatic tail confers lipid solubility on these compounds, which are recruited to cell membranes and become active in various biological pathways. These lipophilic compounds play essential roles in all living organisms (Nowicka & Kruk, 2010). Ubiquinones and menaquinones function as electron carriers in the cellular respiration chain that generates ATP, and as antioxidants that protect membranes against lipid peroxidation. Prenylated hemes are cofactors of the terminal oxidases in the respiration chain that converts oxygen to water. In photosynthetic organisms, chlorophylls absorb energy from light, and plastoquinones transport electrons during photosynthesis. Vitamin E is a group of potent antioxidants that reduce cell damage in plants (Bonitz, Alva, Saleh, Lupas, & Heide, 2011). Other prenylated compounds serve as components of the mycobacterial cell wall, as characteristic lipids of archaeal membranes, and as secondary metabolites. The prenylation reaction that generates these compounds is generally an evolutionarily conserved and rate-limiting step of the biosynthetic pathway (Heide, 2009). The prenyltransferases require two substrates, the prenyl donor and the acceptor. The donor substrate is usually isoprenyl diphosphate (XPP, X stands for the various lengths of the isoprenyl chain). Cleavage of the diphosphate group from the XPP substrate generates a reactive

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carbocation intermediate at the end of the isoprenyl chain. This carbocation reacts with the acceptor substrate to form either a C–C bond or a C–O bond to complete the prenylation reaction.

2. BIOLOGICAL FUNCTION AND ENZYMATIC ACTIVITY OF THE SUBFAMILIES As the prototype of the superfamily, the UbiA enzyme from Escherichia coli is well characterized in biochemistry. UbiA catalyzes the condensation of p-hydroxybenzoate (PHB) with XPP (Fig. 1A). The UbiA activity requires divalent metal ions, preferably magnesium (Melzer & Heide, 1994; Young, Leppik, Hamilton, & Gibson, 1972), which are coordinated by two Asp-rich motifs in UbiA to engage the pyrophosphate group of XPP (Cheng & Li, 2014). The length of XPP can vary from 2 to 10 isoprenyl units (each unit contains five carbons or C5). The E. coli UbiA enzyme appears to have low substrate affinity; the Km values for the reaction between PHB and geranyldiphosphate (GPP; C10) are 0.2 and 0.25 mM, respectively. A higher binding affinity is observed for longer substrates, farnesyldiphosphate (FPP; C15) and solanesyl diphosphate (SPP; C45), albeit with a lower enzymatic activity. Despite the promiscuity of XPP chain length, the prenylation occurs only at the meta-position of PHB (Wessjohann & Sontag, 1996). The control mechanism of this regiospecific reaction remains unknown. COQ2, the eukaryotic homolog of UbiA, has a very similar activity to that of PHB prenyltransferase. In fact, COQ2 or UbiA from different organisms can complement the growth of UbiA-deficient E. coli or COQ2deficient yeast (Boehm, Sommer, Severin, Li, & Heide, 2000; Forsgren et al., 2004; Ohara, Kokado, Yamamoto, Sato, & Yazaki, 2004; Ohara, Muroya, Fukushima, & Yazaki, 2009; Ohara, Yamamoto, Hamamoto, Sasaki, & Yazaki, 2006; Okada et al., 2004; Suzuki et al., 1994; Uchida et al., 2000). These results show that both UbiA and COQ2 are indiscriminate with respect to the chain length of XPPs, which are used in these organisms to produce ubiquinones of 6–10 isoprenyl units (Kainou et al., 2001; Meganathan, 2001; Okada, Kainou, Matsuda, & Kawamukai, 1998; Okada et al., 1996, 1997). Clinically, COQ2 has been linked to infantile multisystem disease, in which COQ2 mutations result in primary ubiquinone deficiency (Quinzii, Hirano, & DiMauro, 2007; Quinzii, Lo´pez, Naini, DiMauro, & Hirano, 2008). The MenA enzyme synthesizes menaquinones (Shineberg & Young, 1976; Suvarna, Stevenson, Meganathan, & Hudspeth, 1998; Young, 1975), the quinones most often used by microbes in their respiration chains.

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A COOH

COOH

COOH

1

PP

2

+

3

n

2

UbiA

+ n

3

COQ2

Cleave

4

Carbocation

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H

4

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3

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PHB

n

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B OH

OH

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+

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DHNA

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K3

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OH

OH

HOOC

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PP 3

OH Cleave

H

(or HGGT, HST)

3

OH

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PPP (or GGPP, SPP)

HGA

E DPPR synthase

Cleave

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cis

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O

+

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P 8

HO

cis

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8

trans HO

OH

pRpp

trans

OH

DP

DPPR

F P

P

OH

+

DGGGP synthase

PP 4

O

H

4

H

O

O 4

GGGP

H

4

GGPP

H

DGGGP

Fig. 1 Different reactions catalyzed by UbiA superfamily prenyltransferases (A–H). Unique chemical groups are colored in red and cleavage reactions are indicated by arrows. Abbreviations: DGGGP, digeranylgeranylglycerylphosphate; DP, decaprenyl phosphate; DPPR, decaprenylphosphoryl-5-phosphoribose; GGGP, geranylgeranylglyceryl phosphate; GGPP, geranylgeranyl diphosphate; HGA, homogentisic acid; PPP, phytyl diphosphate; pRpp, phosphoribose diphosphate.

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Fe

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FPP COOH

COOH

COOH

COOH

Heme B

Heme O

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+

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Chlorophyll synthase

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N

N

N

N

PP 3

N

H O

PPP (or GGPP)

N

C

O COOCH3

O

OH

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Chlorophyllide a

H

Chlorophyll a

Fig. 1—Cont’d

Compared to UbiA, MenA recognizes a different aromatic substrate, 1,4dihydroxy-2-naphthoic acid (DHNA). Decarboxylation of DHNA, which removes the 2-carboxyl group, is coupled with the condensation reaction at the same position (Fig. 1B). Recent studies of MenA (Debnath et al., 2012; Dhiman et al., 2009; Kurosu & Crick, 2009; Kurosu, Narayanasamy, Biswas, Dhiman, & Crick, 2007; Li et al., 2014), focused on developing new tuberculosis drugs, have observed that MenA inhibitors showed significant and specific activity against the mycobacterium at the nonreplicating stage (Debnath et al., 2012). UBIAD1 is a prenyltransferase, found in animals, that is used for the biosynthesis of menaquinone-4 (MK4; vitamin K2), which is converted from phylloquinone (K1) obtained from food sources (Nakagawa et al., 2010). UBIAD1 prenylates menadione (K3), which is generated by the removal of the phytyl tail of K1 (Fig. 1C), by either UBIAD1 or an unknown enzyme (Nakagawa et al., 2010). UBIAD1 is involved in maintaining vascular homeostasis (Hegarty, Yang, & Chi, 2013), preventing oxidative damage in cardiovascular tissues (Mugoni et al., 2013), and sustaining mitochondrial function (Vos et al., 2012). Mutations of UBIAD1 in humans cause

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Schnyder crystalline corneal dystrophy (Orr et al., 2007; Weiss et al., 2007) and urologic cancers (Fredericks et al., 2011). Homogentisate prenyltransferases decarboxylate homogentisic acid and attach different tails. Enzymes in this group, which include homogentisate solanesyltransferases (HST), phytyltransferases (HPT), and geranylgeranyltransferases (HGGT), differ, as indicated by their names, by their specificity requirements for the prenyl donor (Fig. 1D). HST, HPT, and HGGT participate in the biosynthesis of plastoquinones, tocopherols, and tocotrienols, respectively (Soll, Kemmerling, & Schultz, 1980), with the latter two groups of compounds collectively known as vitamin E. To improve the vitamin E content in transgenic plants, HPT and HGGT have been extensively explored for metabolic engineering (Cahoon et al., 2003; Collakova & DellaPenna, 2003; Kinney, 2006; Lee et al., 2007; Savidge et al., 2002; Schledz, Seidler, Beyer, & Neuhaus, 2001; Valentin & Qi, 2005; Venkatesh et al., 2006). The DPPR synthase generates a precursor for arabinogalactans, which form the central layer of a highly impermeable cell wall that is required for the survival of Mycobacterium tuberculosis. DPPR synthase is a promising drug target because both the prenyl donor and the acceptor substrates, decaprenyl phosphate (with several cis-bonds) and 5-phosphoribosyl 1-pyrophosphate (Fig. 1E), are not found in humans (Huang et al., 2005, 2008). The catalysis of DPPR synthase is also unique because the prenyl acceptor contains the pyrophosphate; after its cleavage, the resulting carbocation on the ribose reacts to a monophosphate on the prenyl chain. In contrast, for all other enzymes in this superfamily, a carbocation intermediate is generated from cleaving XPP, the donor substrate (Fig. 1). DGGGP synthase generates the skeleton of diether lipids (Zhang & Poulter, 1993) in an unsaturated form (Fig. 1F), which is the common precursor to most diether and tetraether lipids. These lipids form the core structure of unique archaeal membranes, a major evolutionary feature that distinguishes archaea from bacteria and eukaryotes (De Rosa & Gambacorta, 1988; Koga & Morii, 2007). Although most members of the superfamily use acceptors with ring structures (Fig. 1), the prenyl acceptor in DGGGP synthase is a linear compound (Hemmi, Shibuya, Takahashi, Nakayama, & Nishino, 2004; Zhang & Poulter, 1993). Another difference is that the prenylation by DGGGP synthase generates a C–O link on a glycerol moiety. The heme O synthase, or protoheme IX farnesyltransferase, attaches a farnesyl tail to protoheme IX. These enzymes are also named COX10 in eukaryotes and CyoE and CtaB in E. coli and Bacillus subtilis, respectively

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(Fig. 1G). This universal farnesyltransferase generates heme O, which can be subsequently converted to heme A. These hemes are used in different organisms as the prosthetic group of the terminal heme-copper oxidases in the respiration chain, such as the mitochondrial cytochrome c oxidase (complex IV) that reduces oxygen to water (Hederstedt, 2012). As the terminal enzyme for chlorophyll biosynthesis, chlorophyll synthase esterifies chlorophyllide with phytyl or geranylgeranyl pyrophosphate (Fig. 1H) (Beale, 1999; Oster, Bauer, & R€ udiger, 1997; Schmid, Oster, K€ ogel, Lenz, & R€ udiger, 2001; Willows, 2003). Modified compounds were used to deduce the orientation of the chlorophyllide binding (R€ udiger et al., 2005), which may be similar in chlorophyll synthase and COX10, because both enzymes need to accommodate a large porphyrin ring. Overall, prenyltransferases in the UbiA superfamily are characterized by their different preferences for structurally diverse substrates. This superfamily contains a large number of intramembrane prenyltransferases that are highly diverse in sequence and function.

3. PREDICTING THE FUNCTIONAL ROLE OF INTRAMEMBRANE PRENYLTRANSFERASES BY SEQUENCE CLUSTERING ANALYSIS Sequence clustering provides a useful tool for predicting the function of the large number of uncharacterized superfamily members, which are occasionally misannotated in databases. For example, an archaeal homolog from Archaeoglobus fulgidus, for which a crystal structure has been determined (Huang et al., 2014), is either annotated as bacteriochlorophyll synthase in NCBI or is considered to be a homolog close to MenA and UBIAD1. However, sequence clustering suggests that the function of this homolog is relatively distinct from the functions of known groups of enzymes in the superfamily (Li, 2016). The need for functional prediction is particularly compelling in the field of membrane protein crystallography because it is often necessary to screen a large set of protein homologs to identify the very few that can grow diffracting crystals. In a large screen, activity analysis for all candidate membrane proteins is an intimidating task; sequence clustering offers a simple way to judge the candidate proteins’ functional relevance, which can be verified later if a crystallization hit is obtained.

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The first step in sequence clustering is the selection of representative proteins with well-studied biochemical properties from each major subfamily of known functions (Section 2). These representative proteins, and the protein(s) to be predicted for function, are expanded through a PSI-BLAST search to a number of nonredundant proteins of considerable sequence similarity. Clustering of these sequences is performed based on pairwise sequence similarity, and the function of the target protein is predicted by its relative location on a clustered map. The clustering analysis is more accurate than conventional methods such as multiple sequence alignment and phylogenetic analysis, which can accumulate errors when analyzing thousands of proteins (Frickey & Lupas, 2004). This is the case for the numerous prenyltransferases in the UbiA superfamily, and therefore, sequence clustering analysis (Fig. 2) is performed in the following steps.

3.1 Selecting Representative Proteins of Each Major Subfamily The following proteins are selected, covering most of the subfamilies of intramembrane prenyltransferases that have been identified to date (Section 2). UbiA (Melzer & Heide, 1994; Young et al., 1972) and MenA (Suvarna et al., 1998) are from E. coli. COQ2 (Ashby, Kutsunai, Ackerman, Tzagoloff, & Edwards, 1992; Forsgren et al., 2004), UBIAD1 (Nakagawa et al., 2010), and COX10 (Kim, Khalimonchuk, Smith, & Winge, 2012) are from Homo sapiens. Plant homologs include chlorophyll synthases (R€ udiger et al., 2005) from Arabidopsis thaliana and HGGT (Schledz et al., 2001) from Hordeum vulgare. Microbial enzymes include DPPR synthase (Huang et al., 2005) from Mycobacterium tuberculosis and DGGGP synthase (Zhang & Poulter, 1993) from Methanocaldococcus jannaschii. Sequence clustering is performed between these representative proteins and the unknown proteins to be identified for function. As examples for this functional prediction, we chose two archaeal homologs from Aeropyrum pernix and A. fulgidus for which crystal structures have been determined (Cheng & Li, 2014; Huang et al., 2014).

3.2 Sequence Clustering Analysis All the following steps are conducted using the Bioinformatics Toolkit (http://toolkit.tuebingen.mpg.de/). This platform contains multiple tools, including the CLANS program (Biegert, Mayer, Remmert, S€ oding, & Lupas, 2006), that are required for clustering analysis. Use of this single platform for all steps avoids the problem of data format conversion.

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A

UbiA superfamily

DPPR synthase TbDPPRS

homogentisate PT HvHGGT

Af

HsCOX10

Ap HsCOQ2

DGGGP synthase Chlorophyll synthase

HL23 UbiA COQ2

UbiAD1

EcMenA

MenA

MjDGGGPS Archaeal UbiA homolog

COX10

B

HsUBIAD1

AtChlG

COQ2 UbiA EcUbiA

HL45

HL67 136

191

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245

249

302 305

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RAAGCVVNDYADRKFDGHVKRTANR...YP----FMK...AMVDRDDD

RGAGCTINDMWDQDYDKKVTRTANR...YP----LMK...AHQDKRDD

COX10

SCAANSINQFFEVPFDSNMNRTKNR...YT----PLK...LREDYSRG

HGGT

NIYVVGLNQLYDIQIDKINKPGLPL...YSIEAPFLR...DIPDVDGD

DGGGPS

CAYGNVINDIFDIEIDRINKPSRPL...YAKKYKKYK...DFEDMEGD

ChlG

TGYTQTINDWYDRDIDAINEPYRPI...YSAPPLKLK...IVNDFKSV

DPPRS

ASAVYLVNDVRDVEADREHPTKRFR...YC---FGLK...RYAELHLA

MenA

QILSNLANDYGDAVKGSDKPDRIGP...YT---VGNR...NLRDINSD

UBIAD1

HGAGNLVNTYYDFSKGIDHKKSDDR...YTGG-IGFK...NTRDMESD

90

74

112 115

119

Fig. 2 Sequence clustering and conserved regions of UbiA superfamily prenyltransferases. (A) Homology clustering. The representative proteins used in PSI-BLAST search are indicated (prefix: At, Arabidopsis thaliana; Ec, Escherichia coli; Hs, Homo sapiens; Hv, Hordeum vulgare; Mj, Methanocaldococcus jannaschii; Tb, Mycobacterium tuberculosis), and their positions (large spheres) are shown within each groups (small dots). However, a cluster cannot be generated for the Af homolog (yellow–green dots) due to its low homology to other proteins. (B) Alignment of conserved sequence motifs. The conserved Asp residues are colored in red, and other conserved residues in blue.

1. Create an account in Bioinformatics Toolkit, which allows the saving of performed tasks for a period of time. The following steps will take a few hours to finish and some jobs often require rerunning; therefore, it is crucial to save the successful jobs. 2. Perform a PSI-BLAST search for each representative sequence (Section 3.1) to generate a number of similar sequences. Click the Search

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tab on top of the webpage, select for PSI-BLAST search, and input a target sequence. Under Databases, select nr70, which stands for the NCBI protein database of nonredundant proteins filtered for 70% maximum sequence identity. This relatively low level of identity is used to cover a considerable range of sequence variations. Under Search Options, input in Descriptions the number of sequences to be generated. For the 11 representative proteins being analyzed here (Section 3.1), generate 150 sequences from each PSI-BLAST search; this is to keep the total number of sequences below 2000, which is the limit of the online CLANS application (step 4). Repeat this search for each representative sequence by choosing the existing PSI-BLAST job and clicking on Submit with Same Parameters. Click on the job number and select Alignment to view the aligned sequences. Copy the sequences from each PSI-BLAST job, and combine into a single text file; it is important to use a plain-text editor such as Notepad or Vi (instead of Word). Input the combined sequences into the CLANS program under the Classification tab. To remain consistent with step 2, select the same options (PSI-BLAST and nr70). Select the job number in CLANS when finished. Click on View in CLANS and Continue, which will create a new job that generates the CLANS file (filename.clans). Download this file, and also download and install the CLANS application named clans.jar (a JAVA program), following the link (ftp://ftp.tuebingen.mpg.de/pub/protevo/CLANS/) on the webpage. Start the clans.jar application, and click File and Load Runs to load the filename.clans file obtained from step 5. Many small dots appear in the application window; each dot represents a homolog sequence. Press Initiate to randomize the location of sequence dots, and then Start Run. Homologs of significant similarity (e.g., all UbiA proteins) should start to form clusters of sequence dots. Click Stop when these clusters have converged to a relatively stable position. Click Show Connections to draw lines between dots. Each of these gray lines shows the PSI-BLAST (Altschul et al., 1997) comparison of two sequences, with darker lines indicating higher similarity (lower E values). Click Show selected for a pop-up window, select similar homologs, and click OK. Choose from the menu Windows and Edit Groups to color these homologs in groups. Refer to the CLANS manual for more information.

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4. ANALYSIS OF ENZYMATIC ACTIVITY Several factors complicate the activity analysis of intramembrane enzymes. For example, some of these intramembrane enzymes can be active only in the lipid bilayer. For intramembrane prenyltransferases, XPPs with long aliphatic chains are less soluble, which can be problematic when supplied to an in vitro reaction system (Melzer & Heide, 1994). It is unclear whether these substrates can efficiently diffuse across the membrane–water interface or partition into detergent micelles to reach the prenyltransferases. For UbiA and COQ2, in vitro activity assays generally use the enzymes in crude microsomal membranes, which provide a lipid bilayer environment. The activity assays often use GPP (C10), a short and artificial substrate, due to its high solubility and high yield of enzymatic product. Long substrates are less investigated, as they are difficult to obtain by chemical synthesis and are often not commercially available. Nevertheless, SPP (C45) dissolved in dimethyl sulfoxide can be used by microsomal UbiA, although the catalysis is inefficient (Vmax
50 lower than GPP) (Melzer & Heide, 1994). For the MenA enzymes, activity analyses have been reported using either microsomes (Huang et al., 2014) or proteins purified in detergents (Li et al., 2014). The protocol here describes an in vitro assay of E. coli UbiA activity in microsomes and a complementation assay using a quinone-deficient E. coli strain. Because the endogenous UbiA and MenA activities of E. coli interfere with the enzymatic assay, we constructed a BL21(DE3) strain with knockout of the menA gene and conditional knockout of the ubiA gene. This ubiAmenA strain maintains T7 RNA polymerase overexpression system, which can be used to produce constantly high levels of wild-type or mutant proteins for activity analysis. With the engineered T7 system in BL21(DE3), pET vectors (Agilent) are used to express proteins for use in both structure determination (Section 5) and biochemical analysis. Therefore, consistency is maintained between these studies, and subcloning to different expression vectors is avoided. For activity analysis, the ubiA–menA– strain is used to prepare microsomes for an in vitro assay free of endogenous UbiA or MenA activity. Moreover, the activity of wild-type and mutant UbiA or MenA can be conveniently evaluated by their ability to complement this quinone-deficient strain, which is otherwise prohibited from normal growth.

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4.1 Generation of a ubiA–menA– BL21(DE3) Strain Quinones are essential electron carriers in the respiration chain of all living organisms. E. coli is a facultative anaerobic bacterium that produces both ubiquinone and menaquinone. The relative levels of these two types of quinones change with different growth conditions of E. coli. Of the total quinones generated by E. coli, under aerobic conditions, 65% is ubiquinone, and 35% is menaquinone; under anaerobic conditions, this changes to 7% ubiquinone and 93% menaquinone (Wallace & Young, 1977). The key enzymes in the biosynthesis of ubiquinone and menaquinone are, respectively, UbiA and MenA; both of these genes need to be disrupted in order to make a quinone-deficient strain that can be used for the complementation assay. We conduct the gene knockout under aerobic conditions. Since the menA gene is nonessential for the aerobic growth of E. coli, we can simply disrupt menA in BL21(DE3) by replacing the wild-type menA with a menA:: kan allele from a JW3901 strain (Baba et al., 2006). The introduced kanamycin resistance is used as the selection marker for the knockout strain and is subsequently removed from the chromosome. Removal of the kanamycin-resistant (KanR) gene generated from the menA knockout is necessary because subsequent transduction of the deficient UbiA also needs KanR as a selection marker. The ubiA conditional knockout is made by introducing a ΔmalF:: kan…ubiA420 allele into the BL21(DE3) menA strain. The ubiA420 allele carries a point mutation in ubiA, which was originally isolated from a quinone-deficient strain, AN384 (Wallace & Young, 1977). The mutant has a negligible UbiA activity that cannot be detected by either the complementation or microsomal activity assays (Sections 4.2 and 4.3). However, when supplied with 1 mM PHB, this mutant strain can make
20% amount of the ubiquinone as the wild-type strain; this quinone level is sufficient for the growth of E. coli cells. Therefore, the transductants carrying ΔmalF:: kan…ubiA420 are selected, as they can grow only with 1 mM PHB. The protocol of generating the ubia–mena– Bl21(DE3) strain is as follows. 1. Strains and phage. BL21(DE3) and Phage P1 vir are from the lab collection. NK319 (ΔmalF::kan…ubiA420) is a gift from Dr. Jon Beckwith. JW3901 with ΔmenA789::kan is from the Keio collection (Baba et al., 2006). 2. Preparation of Phage P1 lysate from donor strains. (1) Grow the donor strain JW3901 in 5-mL rich media (dissolve 10 g tryptone, 5 g yeast extract, and 5 g NaCl in 1 L H2O, adjust to

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pH 7.2, and autoclave). Grow the donor strain NK319 at 37°C in 5-mL rich media with 1 mM PHB (Sigma-Aldrich, Catalog# H3776) to late log phase (OD600 ¼ 0.8). (2) Add 250 μL of 0.5 M CaCl2 to the 5 mL cultures. Continue to grow the cells for 10 min. Mix 1 mL aliquot of the culture with 100 μL serial dilutions of P1 lysate stock (100, 10–1, 10–2, and 10–3). Incubate at 37°C for 20 min without shaking. (3) During the incubation, warm up agar plates prepared with rich media (rich plates) in a 37°C incubator. (4) Add 20 μL 20% glucose, 20 μL 0.5 M CaCl2, and 2 mL melted rich top agar (0.7% agar in rich media) to the mixture of cells and phages. Vortex 10 s and pour onto prewarmed rich plates. Allow the soft agar to solidify. Incubate the plate at 37°C for at least 8 h. (5) Choose the plate with evenly distributed individual plaques. Add 2-mL rich media to the plate and scrape off the top agar. Transfer top agar and liquid media into a 50-mL centrifuge tube. Add 20 μL of chloroform and vortex for 10 s. Centrifuge the tube at 10,000 rpm for 10 min to remove debris. Collect the supernatant. Store the P1 lysates from JW3901 and NK319 at 4°C in dark. 3. P1 vir transduction. (1) Grow the recipient strain BL21(DE3) at 37°C overnight in 5-mL rich media. (2) Spin down 0.5 mL of the culture, discard the supernatant, and resuspend the pellet into 0.5 mL MC solution (100 mM CaCl2 and 10 mM MgCl2). (3) Mix 100 μL of cells with 100 μL of serial dilutions (100, 10–1, and 10–2) of P1 lysate made from JW3901. Incubate at 37°C for 20 min without shaking. (4) Add 200 μL 1 M sodium citrate and 0.5-mL rich media. Shake the tube at 37°C for 1 h. (5) Spin the cells and phage mixture at 13,000 rpm for 5 min on a tabletop centrifuge. Discard the supernatant and resuspend the pellet in 0.2 mL 1 M sodium citrate. (6) Plate on a rich plate containing 30 mg/L kanamycin and 1 mM sodium citrate to select for colonies that carry the menA::kan allele. Incubate the plate overnight at 37°C. (7) Pick three to four colonies from the kanamycin plate and streak onto a new kanamycin plate. Grow at 37°C overnight. Repeat the colony streaking at least twice to purify the colonies.

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(8) Confirm the P1 transduction. PCR-amplify individual colonies from the kanamycin plate with the menA forward primer AAAGGCCCCATTTTTATTGG and reverse primer TTTGTCAGTTATGCTGCCCA. The colony that has the incorporated menA::kan allele gives a 1300 bp PCR product. For the PCR analysis, include colonies from JW3901 and BL21(DE3) as positive and negative controls, respectively. The BL21(DE3) strain carrying the wild-type menA gives a 970 bp PCR product. 4. Excision of the kanamycin resistance cassette. In the menA::kan allele, the KanR gene is flanked by a short direct repeats (FRT sites). This KanR gene can be excised in vivo from the chromosome by a yeast Flp recombinase. A temperature-sensitive plasmid pCP20 (Cherepanov & Wackernagel, 1995), which expresses Flp, is transformed into the transductant strains to release the KanR gene. (1) Prepare chemical competent cells of BL21(DE3) menA following a standard protocol. (2) Mix 1 μL pCP20 plasmid DNA with 50 μL competent cells and incubate on ice for 30 min. Heat-shock the competent cells by incubating the tube at 42°C for 30 s. Move the tube to ice and let it sit for 5 min. Add 1 mL SOC media to the tube and recover the cells by shaking at 30°C for 1 h. Plate 100 μL cells onto rich plate with 100 mg/L ampicillin. Incubate the plate at 30°C overnight. (3) Pick three to four single colonies, restreak them on rich plate, and incubate the plates at 42°C. (4) Pick one colony from each of the restreaked colonies, and patch them on rich plate, ampicillin plate, and kanamycin plate. Incubate the patches at 30°C overnight. Those that grow only on the rich plate, but not on the ampicillin or kanamycin plate, have lost the pCP20 plasmid (ampicillin resistance) and have the kanR flipped out. (5) Confirm the removal of KanR by colony PCR with menA forward and reverse primers. Colonies carrying KanR generate a 1300 bp PCR product, and those lost KanR give a 180 bp PCR product. 5. P1 transduction of ΔmalF::kan…ubiA420 to BL21(DE3) menA. (1) Perform P1 transduction of ΔmalF::kan…ubiA420 to BL21(DE3) menA–, using the same protocol as described in step 3. The ΔmalF::kan is used because malF and ubiA genes are located only 9000 bp apart on the E. coli chromosome, and therefore, the transduction of ΔmalF::kan and ubiA420 is tightly linked.

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(2) To select for the transductants, plate the cells on kanamycin plate supplied with 1 mM PHB and incubate the plate at 37°C overnight. Pick 8–10 colonies and patch them on kanamycin plates supplied with and without 1 mM PHB. The colonies that cannot grow without 1 mM PHB carry the ΔmalF::kan…ubiA420 allele.

4.2 Complementation Assay of UbiA Activity With the ubiA–menA– BL21(DE3) Strain 1. Prepare calcium-competent cells using a standard protocol, but with the addition of PHB. Inoculate a single colony of the BL21(DE3) ubiA–menA– strain in 5 mL LB with 1 mM PHB. Grow overnight at 37°C. Take 1 mL aliquot to inoculate 100 mL of the same media and grow until the OD600 reaches 0.2–0.4. Leave cells on ice for 10 min. Collect cells by centrifugation at 4°C and resuspend the cell pellet in 12.5 mL ice-cold 0.1 M MgCl2. Centrifuge again, resuspend cells in 5-mL ice-cold 0.1 M CaCl2, and leave on ice for 20 min. Centrifuge and resuspend cells on ice in 4 mL 0.1 M CaCl2, 10% glycerol, 1 mM PHB. Flash-freeze in aliquots. 2. Prepare LB ampicillin plates with 1 mM PHB (PHB+ plates) and without PHB (PHB plates). 3. Transform wild-type and mutant UbiA in pET15b into the calciumcompetent BL21(DE3) ubiA–menA– cells. Split the transformed cells in half and spread on PHB– and PHB+ plates for overnight growth. 4. Wild-type E. coli UbiA is able to complement the growth of doubleknockout cells on PHB plates. In contrast, inactive mutants of E. coli UbiA, such as Arg60Ala and Asn67Ala, can grow only on PHB+ plates. This complementation assay is a stringent test of activity; mutants that cannot rescue the growth on plates may still have a low activity, which can be detected by the in vitro microsomal assay described later.

4.3 Assay of Enzymatic Activity in Microsomes Determination of the UbiA prenyltransferase activity has been described before (Br€auer, Brandt, Schulze, Zakharova, & Wessjohann, 2008; Melzer & Heide, 1994). For each mutant or wild-type E. coli UbiA, a 100 μL reaction is performed. We generally start with 1 mM GPP (Echelon Biosciences) and 1 mM PHB, which is at a high concentration to ensure that, for mutants of low activity, the reaction product is detectable

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by monitoring the UV absorption on HPLC (Section 4.3.3). For the analysis of enzymatic kinetics, different concentrations of GPP and PHB need to be tested. The typical concentration range of these substrates is 0.01–10 mM for E. coli UbiA. 4.3.1 Preparing Microsomes From the Bl21(DE3) ubiA–menA– Strain 1. Transform wild-type or mutant UbiA into the double-knockout cell. Grow the cells on PHB+ plates overnight. 2. Transfer all colonies on the plate to 1 L LB media supplemented with 1 mM PHB. Grow the cells at 37°C in a 2-L flask, shaking at 225 rpm. When OD600 reaches 0.4–0.6, induce protein expression with 0.4–1 mM IPTG for 3 h. 3. Perform all subsequent steps at 4°C or on ice. Collect cells by centrifugation at 4000 rpm in a JS 4.0 rotor (Beckman). 4. Resuspend cells in 20 mL of 50 mM Tris–HCl, pH 8.0, 0.1 M NaCl. Lyse cells immediately by sonication in the presence of the protease inhibitors, 1 mM phenylmethylsulfonyl fluoride (PMSF; Sigma) and 2 mM benzamidine hydrochloride (Sigma). 5. Centrifuge at 5000 rpm for 10 min to remove unbroken cells. 6. Collect membrane fraction by ultracentrifugation at 50,000 rpm for 15 min in a TLA-100 rotor. 7. Resuspend the membrane fraction in 50 mM Tris–HCl, pH 8.0, and 10 mM dithiothreitol (DTT). Centrifuge again to collect the membrane. Repeat the wash by resuspend the membrane in the same buffer and centrifuge. 8. Use a small glass douncer to thoroughly resuspend the pellet in 4 mL of the same buffer. Flash-freeze aliquots with liquid nitrogen. The concentration of wild-type and mutant UbiA proteins in the microsome resuspension can be matched on western blot, then quantified by comparing with dilutions of purified protein (Section 5.3) using an anti-His antibody (Genscript). 4.3.2 Reaction Setup 1. Prepare the following mixture for a 100 μL reaction. Multiply the volumes by the number of samples to be analyzed. 50 μL 100 mM Tris 7.5 10 μL DMSO 0.5 μL 1 M DTT 0.4 μL 1 M MgCl2

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1 μL 100 mM GPP 1 μL 100 mM PHB 2. Mix a 60 μL aliquot of this reaction mixture with 40 μL microsome, prepared as described in Section 4.3.1. Incubate for 2 h at 37°C with nutation. At the end of the reaction, add 1 μL 25 mM 4-phenylphenol as internal control for the subsequent solvent extraction and HPLC analysis. 3. Immediately mix the reaction mixture with 300 μL ethyl acetate. Vortex at top speed for 1 min to extract the product, 3-geranyl-4hydroxybenzoate. Carefully take 200 μL of the top layer and transfer to a new tube. Dry the extraction by centrifuge under vacuum or by opening the tube to air overnight. 4.3.3 HPLC Analysis of Enzymatic Activity 1. Dissolve the dried product in 100 μL methanol and pass through a 0.1 μm centrifuge filter. 2. Transfer the filtered sample to an HPLC vial. Load the samples on an HPLC (Hewlett-Packard) equipped with an autosampler and a mass spectrometer (Agilent 1100). 3. Inject 10 μL sample on the HPLC with a C18 reverse-phase column (Proto 300 C18 5 μm, 250  4.6 mm) preequilibrated in 5% acetonitrile. Run a gradient of 5–95% acetonitrile in 10 min at 1 mL/min and hold for another 2 min at 95% acetonitrile. The geranylated PHB elutes at
5.16 min and the internal control 4-phenylphenol elutes at
4.70 min (Fig. 3A). Quantify the product by peak areas on HPLC and normalize with the internal control. 4. Confirm the reaction product by mass spectrometry (Fig. 3B). The geranylated PHB has an m/z of 275.3. 5. Analyze the enzyme kinetics from multiple samples using the Prism6 software (GraphPad).

5. STRUCTURAL STUDIES OF THE UbiA SUPERFAMILY 5.1 Cloning, Expression, and Selection of Protein Constructs 5.1.1 Cloning Most of the homologs that we have tested in this superfamily are of microbial origin, for which heterogeneous expression in E. coli is the primary choice. The genes of these homologs were either PCR amplified from the genomic

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Fig. 3 Activity analysis of wild-type EcUbiA prepared in microsomes. (A) HPLC profile (254 nm) shows the product, 3-geranyl-4-hydroxybenzoate (G-PHB), and the internal control, 4-phenylphenol (4-PP). (B) Confirmation of the G-PHB peak (from A) by mass spectrometry.

DNA, if available from ATCC, or obtained from gene synthesis services. The genes were subcloned into either the pET15b or the pET20b vector, which adds either an N- or C-terminal His tag to the protein, respectively. 5.1.2 Selection for Protein Expression Level We screen the expression levels of recombinant homolog proteins for the combination of N- or C-His tags, at 37°C or 22°C, and in three E. coli strains, BL21(DE3), C41(DE3), and C43(DE3). The C41(DE3) and C43 (DE3) strains carry mutations on the T7 RNA polymerase (Miroux & Walker, 1996) that hinder the protein expression; slowly expressed proteins

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tend to fold better in membranes, presumably because the folding machinery in E. coli cells is less overwhelmed. To slow down the protein expression, we also perform expression tests at a lower temperature, i.e., 22°C (sometimes 16°C). Western blot toward the His-affinity tag is used as a quick readout for the protein expression levels. 1. Transform different homologs with N- or C- His tags into BL21(DE3), C41(DE3), and C43(DE3). Grow each of these constructs on an LB ampicillin plate at 37°C overnight. 2. Inoculate a single colony from each plate to 5 mL LB ampicillin media and grow at 37°C. Add 1 mM IPTG at OD600 0.6–0.8. Immediately split 2.5 mL culture to another tube and induce at 22°C overnight. Keep the rest of the 2.5 mL culture at 37°C and induce for 3–6 h. 3. Centrifuge to collect 50–100 μL of cells at OD600 ¼ 1 or an equivalent volume. Match the volume of cell samples by their cell density (OD600) to ensure that the same number of cells are compared for expression levels. 4. Resuspend the cell pellet with 50 μL 2  SDS-PAGE sample buffer. Apply to a 12% SDS-PAGE with 1.5-mm thick wells. Heating the cells at 95°C in sample buffer gives a less viscous solution, but may change the gel mobility of some membrane proteins. Therefore, we generally skip the heating step (or heat at 60°C in some cases), but use a small amount of cells to lower the viscosity. 5. Detect the expression level of His-tagged protein by western blot. Transfer protein bands on the SDS-PAGE gel to a nitrocellulose membrane at 100 V for 1 h using a wet-blotting system (BIORAD). Block the membrane at 22°C for 1 h in 10 mL TBS-T (50 mM Tris, pH 7.4, 150 mM NaCl, and 0.1% (v/v) Tween 20) with 5% milk. Incubate the membrane at 22°C for 1 h with 5 μL mouse anti-His antibody (Genscript) in 10 mL TBS-T. Wash the membrane three times with 10 mL TBS-T for 10 min each time. Incubate the membrane at 22°C for 1 h with 2 μL of goat antimouse IgG peroxidase (Sigma) in 10 mL of TBS-T. Wash the membrane three times with 10 mL of fresh TBS-T for 10 min. 6. Immediately add ECL western blotting substrate (Pierce). Develop the film in a short exposure time (10–30 s). A strong band on the film indicates a well-expressed protein (Fig. 4A). Constructs showing weak bands are difficult to pursue further because the protein usually cannot be purified to crystallographic quality in the end; the purification protocol described here relies primarily on an effective His-affinity chromatography step (Section 5.3).

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5.1.3 Selection for Properly Folded Proteins This first round of western blot analysis (Section 5.1.2) significantly narrows down the protein targets: only those that are sufficiently expressed are pursued further (Fig. 4A). However, this preliminary test cannot exclude misfolded and aggregated proteins, such as those in the inclusion body. To overcome this problem, we take the assumption (Sonoda et al., 2010) that proteins that are properly folded in membranes can be efficiently extracted by n-dodecyl-β-D-maltopyranoside (DDM), the most commonly used detergent in membrane protein crystallography.

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1. Freshly transform selected constructs with high level of protein expression. Grow and induce cells with the best conditions identified earlier. 2. Resuspend and sonicate the same amount of cells (matched by OD600) from
10 mL culture in 500 μL of 20 mM Tris–HCl 7.5, 0.1 M NaCl. Add powder of DDM (Anatrace, Sol-Grade) to 1% final concentration, and incubate for 1 h with nutation for membrane protein extraction. For comparison, save an aliquot of cells without DDM extraction. 3. Centrifuge and run the supernatant from DDM extraction on western blot. Run the unextracted cells side by side. The protein constructs that can be effectively solubilized by DDM are pursued further. 5.1.4 Selection for Monodispersed Proteins For each selected protein, a medium scale of purification is conducted from a 2 L culture. We generally purify six proteins (i.e., a total of 12 L culture) at the same time, following a protocol described in Section 5.3. The proteins purified after Ni chromatography can be directly compared on SDS-PAGE for amount and purity. Proteins to be pursued further should be relatively pure at this stage (Fig. 4B), and the amount purified from the 2 L culture is generally sufficient for the subsequent analysis of the elution profile by size exclusion chromatography. Protein constructs that give sharp and symmetrical peaks at the proper elution volume (Fig. 4C) are selected for large-scale purification and crystallization trials (Sections 5.2–5.4). 5.1.5 Results and Other Considerations An alternative approach, which is more efficient than the western blot method described here, is to clone proteins with GFP tags and use fluorescence size exclusion chromatography (FSEC) to screen for the expression level and elution profile at the same time (Kawate & Gouaux, 2006). The FSEC method requires cloning into the vectors with the GFP sequence engineered, in addition to cloning into the pET vectors. Conversely, if the GFP vectors are directly used for expression and purification, cleavage of the GFP tag is normally required at a later step. In some cases, although the GFP-tagged protein gives a sharp and symmetrical peak at the initial FSEC test, removing the GFP tag by protease digestion may destabilize the target protein and lead to aggregation. Although our lab routinely uses FSEC for membrane proteins expressed in eukaryotic systems, for E. coli expression, our experience is that the western blot method is as effective as FSEC in selecting the constructs suitable for crystallographic studies.

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After screening large numbers of UbiA and MenA homologs, we identified an archaeal UbiA homolog from A. pernix (ApUbiA) that is well expressed and shows a good profile on size exclusion chromatography (Fig. 4C). Interestingly, one of the few homologs that behave well in our hands, the homolog from A. fulgidus (Af ), was also independently identified by another group (Huang et al., 2014); this homolog was initially found during a structural genomics search and also succeeded in structure determination. This coincidence suggests that there are only a small number of protein homologs in this large superfamily that behave well enough to be readily pursued for crystallographic studies. Other protein homologs in this family remain possible targets for structural determination, but to be successful, these may require more optimization efforts to find better expression conditions, suitable detergents, and proper protein constructs, such as truncations that remove flexible or degradable regions. The ApUbiA homolog is best expressed in C43(DE3) cells at 22°C. In addition, minimal media gives a significantly higher level of protein expression compared to LB media. This growth condition was found during the process of crystal structure determination, in which the minimal media is routinely used for selenomethionine (SeMet) labeling to obtain experimental phasing. The improvement of protein expression in SeMet media has also been occasionally observed for other membrane proteins (Zimmer, Nam, & Rapoport, 2008). Therefore, SeMet labeling should probably be always tested for new membrane proteins under investigation, with the improvement of expression level as a bonus to obtaining experimental phasing. The following protocol describes the expression and purification of the ApUbiA protein with SeMet labeling, which can serve as a model for producing other microbial proteins in the UbiA superfamily for crystallographic studies.

5.2 Expression of the ApUbiA Protein in a Minimal Media Used for SeMet Labeling The following steps are modified from a published protocol (Doublie, 2007). Six amino acids (Lys, Thr, Phe, Leu, Ile, and Val) are added before induction to inhibit Met biosynthesis, which in turn allows efficient SeMet incorporation. 1. Prepare the following. (1) 1.5 L of 10 M9 medium. Dissolve 132.9 g Na2HPO47H2O, 58.5 g KH2PO4, 9.75 g NaCl, and 19.5 g NH4Cl in 1.5 L H2O and transfer to a 2-L flask.

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(2) For 1 L adaptive media (below), add 50 g glucose and 5 g LB to 920 mL H2O in a 2-L flask. (3) For 12 1 L expression media (below), add 800 mL H2O and 50 g glucose to each 2-L flask. (4) Autoclave media (1) to (3), along with empty centrifuge bottles and empty cylinders (used below). (5) Dissolve 65 g yeast nitrogen base (YNB; Sigma) in 1 L H2O and sterile filter (10 6.5% YNB stock solution). (6) Make 200 mL of 10 M NaOH. (7) After the media is cooled down, use the sterilized cylinder to (a) add 80 mL of 10 M9 medium to make 1 L adaptive media. (b) add 100 mL of 10  M9 medium and 100 mL of 10 YNB to make each 1 L expression media. 2. Grow 12 L cells to express the ApUbiA protein with SeMet labeling. (1) Transform 1 μL plasmid DNA, pET15b-ApUbiA, to 30 μL calcium-competent C43(DE3) cells. Grow on LB ampicillin plate overnight. (2) Scrape all colonies on the plate into the 1 L adaptive media. Grow at 37°C until OD600 reaches 0.6–0.8. (3) Collect cells by centrifuge at 4000 rpm at 4°C for 10 min. Resuspend cells in 240 mL of expression media. Add 20 mL of cells to each 1 L of expression media in the 12 flasks. Grow at 37°C until OD600 reaches 1.0. (4) Add 4.5 mL 10 M NaOH dropwise to each 1 L of expression media. Change temperature to between 16°C and 22°C. After 20 min, add seven amino acids: Lys, Thr, and Phe at 100 mg/L; Leu, Ile, and Val (all from Sigma) at 50 mg/L; and SeMet (Anatrace) at 60 mg/L. Resuspend the amino acids by stirring in 120 mL H2O first, and pipet 10 mL of the suspension to each 1 L expression media. (5) After 30 min, add 0.4–1 mM IPTG to induce protein expression. Shake at 180 rpm for 13–17 h. (6) Centrifuge at 4000 rpm at 4°C for 20 min to collect the cells. The cells can be frozen by liquid nitrogen and stored at –80°C for later use.

5.3 Protein Purification 1. Perform all subsequent steps in this section at 4°C or on ice. Thaw and resuspend the cells from 12 L culture (Section 5.2) in 120 mL buffer

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A containing 20 mM Tris–HCl, pH 8.0, 0.3 M NaCl, and 10% glycerol. Use a food processor to thoroughly resuspend the cells. Lyse the cells by passing twice through a microfluidizer (Microfluidics M-110L) with the pressure set at 15,000 psi. After the first pass, add 1.2 mL of 100  stock solution of protease inhibitors, which is made by dissolving 0.1 M PMSF and 0.2 M benzamidine in ethanol. Centrifuge the lysed cells at 5000  g for 10 min to remove unbroken cells. Pour the supernatant to 2 of 70 mL ultracentrifuge tubes and collect the cell membrane by ultracentrifugation at 42,000 rpm in a Ti45 rotor (Beckman) for 1 h. Remove supernatant and use a 55-mL glass douncer (attached to an electrical overhead stirrer) to homogenize the membrane in a final volume of 60 mL buffer A with 1% (w/v) DDM (Sol-Grade from Anatrace). Stir the mixture for 1 h to thoroughly solubilize the membrane in DDM. Ultracentrifuge at 42,000 rpm for 30 min in a Ti45 rotor. Prepare a 10 mL suspension of Ni-NTA resin (Qiagen) in a 50-mL gravity-flow column. Wash the resin with 100 mL water in steps to remove ethanol. To equilibrate the resin, wash with 100 mL Buffer A in 10 mL steps, and then 10 mL Buffer A with 0.03% DDM. Resuspend the resin by 5 mL Buffer A with 0.03% DDM. Add the resin to the supernatant from step 6. Stir the Ni-NTA resin with the DDM solubilized protein for 1 h for batch binding. Collect the resin back in the 50 mL column. Carry out all subsequent washing and elution in 5–10 mL additions at a time. Thoroughly wash the resin in the following steps: 300 mL Buffer A with 0.03% DDM; 300 mL Buffer A with 0.03% DDM and 20 mM imidazole; 600 mL Buffer A with 0.06% 6-cyclohexyl-1-pentyl-β-D-maltoside (CYMAL6; Anatrace, Sol-Grade) and 40 mM imidazole. Elute the protein in 40 mL Buffer A with 0.06% CYMAL6 and 0.3 M imidazole. Run an SDS-PAGE gel of aliquots from each of the wash and elution steps. The samples are mixed with SDS-PAGE sample buffer without heating. Proceed only if a significant amount of protein is purified in the elution fraction. Concentrate the eluted protein to 0.5 mL with a 50-kDa cutoff Amicon Ultra (Millipore) centrifuged at 2000–4000 rpm. Frequently check and mix the sample to avoid overconcentrating. Centrifuge in a 0.01 μm Ultrafree filter unit (Millipore) to clear up any precipitation in the sample.

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12. Immediately apply the filtered sample on a Superdex 200 size exclusion column (GE Healthcare) that has been preequilibrated in a buffer containing 10 mM Tris–HCl, pH 8.0, 0.1 M NaCl, and 0.25% 5-cyclohexyl-1-pentyl-β-D-maltoside (CYMAL5; Anatrace, SolGrade). Collect fractions and run SDS-PAGE gel. 13. Concentrate the protein to 17.5 mg/mL. Determine the protein concentration by OD280 using a Nanodrop spectrophotometer.

5.4 Selection of Detergents Careful detergent selection is essential for work with membrane proteins, because their structural stability strongly depends on the detergent being used. Moreover, membrane protein crystals are formed both by the protein and by associated detergent micelles; therefore, the micelle size is an important factor in improving crystal diffraction. The choice of detergents is essential in crystallizing the ApUbiA protein and improving crystal diffraction. This protein can be crystallized in DDM and in other detergents that have a maltose head group and a similar length aliphatic chain. Compared to crystallization in DDM, crystallization ˚ , which allowed in CYMAL5 improved the diffraction from
8 to 3.3 A the building of a reliable structural model. In addition, ApUbiA that is purified in the lauryldimethylamine-N-oxide (LDAO) detergent, which has a relatively small micelle, can be crystallized in a different crystal form for which the diffraction spots beyond 3 A˚ can be observed. However, the LDAO crystal form is severely twinned and was not able to provide any more structural information than the 3.3 A˚ model of the CYMAL5 crystals. The initial screen of the detergents is usually conducted at the gel filtration step. The ApUbiA protein is purified in DDM until its elution from the Ni-NTA column, with the assumption that DDM is a mild detergent that maintains stability for most of the membrane proteins (Sonoda et al., 2010). The protein is concentrated in the buffer with DDM (Section 5.3) and applied to the Superdex-200 column that has been preequilibrated with the buffer containing various detergents. The detergents with the maltose head group and varying aliphatic groups, such as n-decyl-β-D-maltoside (DM) and CYMALs, are generally tested first. In addition, we chose detergents that have been reported to be most successful in crystallization. As nicely summarized by previous reviews (Newstead, Kim, von Heijne, Iwata, & Drew, 2007; Sonoda et al., 2010, 2011), these detergents include

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the glucosides, octyl- (OG) and nonyl-glucopyranoside (NG); octaethylene glycol monododecyl ether (C12E8); LDAO; and more recently, detergents with two maltose head groups, such as lauryl maltose neopentyl glycol (LMNG). If a detergent other than DDM gives a sharp and symmetrical peak at the gel filtration step, this detergent can be further tested for use at an earlier step in purification, such as the initial solubilization step or the wash and elution step after the protein has bound to the Ni-NTA resin. For ApUbiA, we found that the best protocol is to solubilize the protein in DDM, wash and elute with CYMAL6 on Ni-resin, and run gel filtration in CY MAL5 (Section 5.5).

5.5 Crystallization and Structure Determination Structures of two archaeal homologs have been determined in the UbiA superfamily. The ApUbiA homolog was crystallized in detergent and the ˚ (Cheng & Li, 2014). A 3.6 A ˚ structure was apo crystal diffracted to 3.3 A determined with geranyl thiolodiphosphate (GSPP); this noncleavable substrate was used because UbiA was previously reported to hydrolyze GPP even in the absence of PHB (Br€auer et al., 2008). The crystals were also soaked with PHB and Mg, both of which were modeled into the structure at this moderate resolution. The Af homolog was crystallized in liquid cubic phase (LCP) and the structures with GPP or dimethylallyl diphosphate (DMAPP; C5) were determined to be 2.4–2.5 A˚ (Huang et al., 2014). The positions of Mg2+ ions in this structure were confirmed by anomalous signals from Cd, a heavier divalent atom in substitution for Mg. Crystals grown in LCP generally diffract better than those grown in detergents because the protein molecules are more closely packed with interactions by membrane domains. Besides the Af homolog, we have recently obtained LCP crystals of other proteins in the superfamily (unpublished data). As crystallization in detergent is a more straightforward method, the protocols described later put more emphasis on the LCP method. 5.5.1 Crystallization in Detergents Initial crystallization screens of ApUbiA are performed with chemical solutions from commercial screening kits (Hampton Research and Qiagen), with the aid of a Mosquito crystallization robot (TTP Labtech). Crystallization conditions are optimized for buffer conditions, protein concentrations, and additives. The protein in LDAO crystallizes at a concentration of

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13 mg/mL with an initial 1:1 mix of the protein to 0.12 M Na2SO4, 20.5% (w/v) PEG400, and 0.1 M Tris–HCl, at pH 8.0. The crystals are grown by the hanging drop vapor diffusion method at 22°C, and these hexagonal plates reach final size in 2 weeks. The protein in CYMAL5 crystallizes at 17.5 mg/mL with 0.1 M Na2SO4, 26.5% (w/v) PEG400, and 0.1 M sodium cacodylate, at pH 5.0. The crystals are rod-shaped and grow to final size in 4 weeks. The crystals are soaked in 5 mM GSPP, 10 mM MgCl2, and 10 mM PHB for 1 h. Both the native and soaked crystals are directly flash-frozen in liquid nitrogen. 5.5.2 Crystallization in LCP The LCP method has been described before (Liu & Cherezov, 2011); the following protocol is adapted to the UbiA superfamily proteins. The LCP lipid we normally use is monoacylglycerol (MAG9.9) monoolein, and shortchain MAGs can be explored to improve crystal diffraction. 1. Reconstitute the protein in LCP. (1) Purify a UbiA superfamily protein in a detergent solution. Concentrate the protein to
30 mg/mL. Avoid overconcentrating the detergent during this process. To do this, choose and combine only the peak fractions from the size exclusion chromatography and concentrate the complex of protein (usually
30 kDa) and detergent micelle in an Amicon Ultra Centrifugal Filter with a 50-kDa cutoff. (2) Warm up a 70 μL monoolein aliquot (NU CHEK) at 40°C for few minutes, until the lipid melts. (3) Load one of the LCP mixing syringes (Art Robbins) with the melted monoolein. Remove the plunger from the syringe. Use a 200 μL pipette mounted with a yellow pipet tip to add the lipid from the top of the syringe. This tip fills directly in the syringe, and the lipid can be pushed slowly down with the syringe slightly tilted. After adding the monoolein, put the plunger into the syringe. If air bubbles are trapped, push the plunger down slowly and then move it up quickly a few times, which generally removes the bubbles. Record the volume of the lipid in the syringe. (4) Load another syringe with the protein solution through the syringe tip. The volume of the protein is generally 2/3 of the lipid volume. Take caution to avoid generating bubbles in the protein solution. (5) Connect both syringes together through a syringe coupler. (6) Push the two syringe plungers back and forth slowly to mix the lipid and protein through the needle part of the coupler. This mixing

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usually takes more than 100 times. The protein and lipid mixture is initially a white color, and it becomes transparent after thorough mixing. This indicates that a homogeneous lipid mesophase has been obtained through mechanical mixing, and that the protein has been reconstituted in the lipid bilayer of LCP. (7) Push all mixture to one syringe. Take off the other syringe and the coupler. Add on a needle and a metal ring to the syringe in use. 2. Set up LCP crystallization with a Gryphon LCP robot (Art Robbins). (1) Clamp the syringe in position on the dispensing arm of the robot. Place an LCP plate (molecular dimensions or custom-made glass plates) and a 96-well block containing the precipitant solutions on the designated decks of the robot. (2) Load the 96-tip head of the robot with precipitant solutions. Commercial screening kits, such as MB Class Suite and MB Class Suite II (Qiagen), should be diluted to 70% to lower the precipitant concentration because high precipitant concentration may disrupt the LCP mesophase (Johansson, W€ ohri, Katona, Engstr€ om, & Neutze, 2009). (3) Dispense the LCP mixture and precipitants with the robot programmed as follows: sequentially dispense 100 nL of the mesophase protein from the syringe onto the 96 wells of LCP plate; dispense 800 nL of precipitant solutions on top of the mesophase with the 96-tip head of the robot. The entire dispensing process takes about 2 min to finish. (4) Remove the LCP plate from the platform of the robot and seal with a cover glass. Ensure that all 96 wells are centered and fully covered. Label and place the plate in incubator (usually at 22°C) for crystal growth. (5) Remove the precipitant block from the deck of the robot, seal it tightly, and put it back into storage. Remove the syringe from the dispensing arm of the robot. Dismantle the syringe and wash the syringe, needle, and ferrule with ethanol. Air-dry these parts for future use. 3. Harvest LCP crystals. (1) Place an LCP plate on the stage of a light microscope. Use a glasscutting tool (Hampton Research) to draw two concentric circles on the cover glass around the well that contains the LCP crystals. Break up the glass along the circles to release the central piece of cover glass. Remove the dust of broken glass with moistened Kimwipes.

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Grip the cover glass with a fine-tipped tweezer, tilt the glass piece away, and take it off the well. (2) The mesophase bolus can be on either the base plate or the cover glass. Zoom in the microscope to get a clear view of the LCP crystals. Use a cryo-loop to take the crystals out from the freshly exposed mesophase lipids. Plunge the crystals into liquid nitrogen to freeze. To avoid crystal damage, harvest the LCP crystals as fast as possible after breaking the cover glass.

5.6 Structure Comparison of Intramembrane and Soluble Prenyltransferases Besides intramembrane prenyltransferases, several classes of soluble enzymes catalyze either the fusion of two molecules, a prenyl donor with an acceptor, or similar fusion reactions within one molecule (Brandt et al., 2009). Central to these reactions are a resonance-stabilized carbocation intermediate that is generated after the diphosphate group of XPPs is cleaved off. Terpene synthases join this carbocation intramolecularly, often with a C–C double bond, to form a large variety of circlized molecules. Isoprenyl diphosphate synthases catalyze the elongation of linear XPPs by the intermolecular reaction of the carbocation from the allylic XPP to the unsaturated terminal of a homoallylic substrate, isopentenyl diphosphate (IPP). Protein prenyltransferases add the carbocation to the sulfur atom of a cysteine residue in a protein. Soluble aromatic prenyltransferases catalyze the carbocation reaction to aromatic compounds, with the formation of C–C, C–O, or C–N bonds. Here, we will focus on the structural comparison of these enzymes to explore their substrate specificity and their different strategies of catalysis. Two structures of intramembrane prenyltransferases have been determined (Fig. 5A). The Ap (Cheng & Li, 2014) and Af homologs (Huang et al., 2014), both of archaeal origin, contain nine-transmembrane helices (TM) that arrange counterclockwise to form a U-shaped architecture. An extramembrane cap domain forms over the membrane domain and contains most of the conserved residues, including the two Asp-rich motifs. The Asp residues coordinate Mg2+ ions to engage the pyrophosphate group of XPP. These well-conserved Asp residues are essential for the activity of various enzymes in the superfamily (Br€auer, Brandt, & Wessjohann, 2004; Cheng & Li, 2014; Huang et al., 2008, 2014; Ohara, Mito, & Yazaki, 2013, Ohara et al., 2009; Saiki, Mogi, Hori, Tsubaki, & Anraku, 1993; Stec & Li, 2012).

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A

Ap

Af Cap domain

Cap domain HL23

HL23

HL67

HL67

HL45 HL45 TM7

TM6

TM4 PHB TM8

TM9 TM5

GPP

Membrane boundary

GSPP

TM6

TM3

TM7 TM3

TM1

TM5

Membrane domain

TM2 TM9

TM8

TM1

TM2

B

TM4

Ap

Af Unrestricted central cavity

Membrane boundary TM8

TM8

TM9

Restricted pocket

TM9

TM1 TM1

Long tunnel

C

FPPS

DMSPP IPP

NphB

UPPS

FSPP P GSPP DHN

Fig. 5 Structural comparison of intramembrane and soluble prenyltransferases. (A) Crystal structures of intramembrane prenyltransferases. Structures of the Ap (PDB: 4OD5) and Af (PDB: 4TQ3) homologs are shown in the same orientation. The cap domains are colored in pink, and TM domains of Ap and Af in cyan and green, respectively. The Mg2+ ions are shown as cyan spheres. (B) Comparison of the Ap and Af substrate-binding pockets. (C) Overall structures of soluble prenyltransferases with substrates. trans-, cis-, and aromatic prenyltransferases are each shown with a representative structure, farnesyl pyrophosphate synthase (FPPS; PDB 1RQI), undecaprenyl pyrophosphate synthase (UPPS; PDB 1X06), and NphB (PDB 1ZB6), respectively.

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D

Ap D182

Af

FPPS

D186

Q241

DMSPP

D202 R63

D58

Mg

Mg

K119

Mg

K146 Y139

N50

D105

R69

Mg

D72

H98

Mg

D175 Y115 GSPP

Mg

Q179 K202 Mg C1

R318

GPP

D54 R67

Y178

D244 T203

D198

K66

S140

R22

C1

PHB

K258

IPP

R116

N68 R117

R43

UPPS

NphB R228

R30 Mg

N28 H43

P D26 C1 N74

R39

Mg

FSPP

K284

D62 K119 C1 D110

R77

Y282 N173

K118

Y216

S51 GSPP DHN Y288

Fig. 5—Cont’d (D) Comparison of the active site residues and Mg positions in all these structures. The carbocation is generated on the C1 atom (red). Abbreviations: DHN, dihydroxynaphthalene, an analog of the aromatic substrate in NphB structure; DMSPP and FSPP, uncleavable thiophosphate derivative of DMAPP and FPP, respectively; P, a phosphate in substitute of IPP in the UPPS structure.

The Ap and Af structures differ significantly in the two substrate pockets that bind the prenyl donor and acceptor (Fig. 5B). ApUbiA has a large central cavity that opens laterally to the lipid bilayer, thereby creating a unique passage to the active site that may facilitate the binding of long-chain XPP substrates and the release of their prenylated products in membranes. This unrestricted binding chamber also explains the chainlength promiscuity of UbiA and COQ2; these enzymes accept a variety of IPP lengths to generate ubiquinones with 6–10 isoprenyl units, which are characteristic of different species. The central cavity contains a hydrophobic bottom wall and a small basic pocket, which have been proposed to bind the isoprenyl chain of XPP and the aromatic substrate, respectively (Cheng & Li, 2014).

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In contrast, the central cavity of the Af homolog is not directly exposed to lipid. Instead, a long empty tunnel comes off the active site, with a small opening to the membrane. This long tunnel is shaped to bind a linear substrate, which is presumably the prenyl acceptor. Conversely, the prenyl donor, GPP or DMAPP, is observed to bind to another pocket in the structure. This pocket is restricted by the protein backbone (Fig. 5B) and therefore is not capable of accommodating longer XPPs. The different substrate-binding pockets in the two structures are generated by the different spacings between TM1, TM8, and TM9. The large separation between TM1 and TM9 in the Ap structure generates the open lateral portal. In contrast, TM1 and TM9 in the Af structure are close together, thereby blocking the direct opening of the Af central cavity to lipids. Because TM9 is positioned away from TM8, a larger space is generated between these helices, thereby creating the long tunnel. The variation between the two structures implies that the relative positions of these three TMs largely define the substrate specificity, and this may be a general mechanism that is used also by other enzymes in the superfamily to determine their substrate preferences. The 9-TM structure of intramembrane prenyltransferases is surprisingly similar to the isoprenyl synthase fold shared by trans-prenyltransferases and terpene synthases (Fig. 5C). A DALI search (Holm & Rosenstr€ om, 2010) indicates that the closest match to intramembrane prenyltransferases is the farnesyl pyrophosphate synthase (Hosfield et al., 2004; Kellogg & Poulter, 1997; Wallrapp et al., 2013), which has seven helices superimposable to the TM 1–7 in ApUbiA (Cheng & Li, 2014). Similar to enzymes in the UbiA superfamily, trans-prenyltransferases and terpene synthases also contain two Asp-rich motifs that bind two or three Mg2+ ions (Fig. 5D). In contrast, cis-prenyltransferases, although they have a similar function as the trans-type, adopt a different structural fold, using a single Asp or Glu residue to coordinate one Mg2+ ion for enzymatic activity (Fig. 5C and D). Soluble aromatic prenyltransferases share a similar function to the intramembrane prenyltransferases, but there is no sequence or structural similarity. All soluble aromatic prenyltransferases have a PT-barrel fold made of tandem ααββ structural repeats (Fig. 5C). The α-helices surround a β-barrel, inside of which are the substrate-binding pockets (Jost et al., 2010; Kuzuyama, Noel, & Richard, 2005; Metzger, Keller, Stevenson, Heide, & Lawson, 2010, Metzger et al., 2009; Saleh, Haagen, Seeger, & Heide, 2009). Soluble aromatic prenyltransferases do not contain Asp-rich motifs, and the divalent metal ion is dispensable in some cases (e.g., indole

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prenyltransferases). Instead, an aromatic side chain in these enzymes may stabilize the carbocation by a cation–π interaction (Fig. 5D). For the prenylation of aromatic compounds, the reactive carbocation needs to be shielded from water (Bonitz et al., 2011; Metzger et al., 2009). In soluble APTs, the shielding is achieved by the β-barrel, whereas in intramembrane prenyltransferases, the TMs and surrounding lipids may play a similar role. Overall, the prenyltransferases known to date show large variation in their structures and catalytic mechanisms. Structural and biochemical analyses of the intramembrane prenyltransferases reveal new strategies of catalysis that have been adapted to the membrane environment. The active site of these enzymes is opened to the lipid to allow ready access to long-chain substrates, as well as the efficient dissociation of prenylated products of long tails. Catalysis is permitted with this lipid opening because the reactive carbocation is protected from water by the surrounding lipids. Future studies, using the methods described in this chapter, will reveal detailed catalytic mechanisms, such as the mechanism of generating and stabilizing the carbocation, and the condensation reaction that joins the prenyl donor and acceptor together.

ACKNOWLEDGMENTS W.L. is supported by an R01 (HL121718) and an R00 Grant (HL097083) from National Heart, Lung, and Blood Institute, a Grant-in-Aid (14GRNT20310017) from American Heart Association, and a scholar award from the American Society of Hematology.

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CHAPTER FOURTEEN

Functional Study of the Vitamin K Cycle Enzymes in Live Cells J.-K. Tie1, D.W. Stafford University of North Carolina at Chapel Hill, Chapel Hill, NC, United States 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 1.1 Functional Study of VKOR by In Vitro Activity Assay 1.2 Function Study of GGCX by In Vitro Activity Assay 1.3 Function Study of VKR by In Vitro Activity Assay 2. Cell-Based Assay for Functional Study of Vitamin K Cycle Enzymes 2.1 Rationale 2.2 Establishment of the FIXgla-PC Reporter Cell Lines 2.3 ELISA-Based Evaluation of the Reporter Protein Carboxylation 2.4 Applications 3. Functional Study of VKOR Using TALENs-Mediated Gene Knockout Reporter Cells 3.1 Rationale 3.2 Establishment of the TALENs-Mediated VKOR/VKORL Knockout Reporter Cell Line 3.3 Functional Study of VKOR Using the DGKO Reporter Cells 3.4 Applications 4. Functional Study of GGCX Using CRISPR-Cas9-Mediated Gene Knockout Reporter Cells 4.1 Rationale 4.2 Establishment of the CRISPR-Cas9-Mediated GGCX Gene Knockout Reporter Cell Line 4.3 Functional Study of GGCX Using the GGCX-Deficient Reporter Cells 4.4 Applications 5. Concluding Remarks References

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Abstract Vitamin K-dependent carboxylation, an essential posttranslational modification catalyzed by gamma-glutamyl carboxylase, is required for the biological functions of proteins that control blood coagulation, vascular calcification, bone metabolism, and other important physiological processes. Concomitant with carboxylation, reduced

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vitamin K (KH2) is oxidized to vitamin K epoxide (KO). KO must be recycled back to KH2 by the enzymes vitamin K epoxide reductase and vitamin K reductase in a pathway known as the vitamin K cycle. Our current knowledge about the enzymes of the vitamin K cycle is mainly based on in vitro studies of each individual enzymes under artificial conditions, which are of limited usefulness in understanding how the complex carboxylation process is carried out in the physiological environment. In this chapter, we review the current in vitro activity assays for vitamin K cycle enzymes. We describe the rationale, establishment, and application of cell-based assays for the functional study of these enzymes in the native cellular milieu. In these cell-based assays, different vitamin K-dependent proteins were designed and stably expressed in mammalian cells as reporter proteins to accommodate the readily used enzyme-linked immunosorbent assay for carboxylation efficiency evaluation. Additionally, recently emerged genomeediting techniques TALENs and CRISPR-Cas9 were used to knock out the endogenous enzymes in the reporter cell lines to eliminate the background. These cell-based assays are easy to scale up for high-throughput screening of inhibitors of vitamin K cycle enzymes and have been successfully used to clarify the genotypes and their clinical phenotypes of enzymes of the vitamin K cycle.

1. INTRODUCTION Vitamin K-dependent (VKD) carboxylation is an essential posttranslational modification that converts certain glutamate (Glu) residues to gamma-carboxyglutamate (Gla) residues in VKD proteins. The modification involves the addition of a carboxyl group to the gamma carbon of Glu residues in the VKD protein. Carboxylation is catalyzed by the enzyme gamma-glutamyl carboxylase (GGCX), which utilizes a reduced form of vitamin K (vitamin K hydroquinone, or KH2), carbon dioxide, and oxygen as cofactors. Concomitant with each Glu modification, KH2 is oxidized to vitamin K 2,3-epoxide (KO). Since vitamin K is rapidly catabolized in the human body (Shearer & Newman, 2014), KO must be reused and is converted back to KH2 through a two-step reduction by the enzymes vitamin K epoxide reductase (VKOR) and the as-yet-unknown vitamin K reductase (VKR) in a pathway known as the vitamin K cycle (Fig. 1). Because of the hydrophobic characteristics of vitamin K, enzymes in the vitamin K cycle are likely integral membrane proteins that reside in the endoplasmic reticulum (ER). VKD carboxylation has been associated mostly with coagulation, because it was originally observed in clotting factors (Nelsestuen, Zytkovicz, & Howard, 1974; Stenflo, Fernlund, Egan, & Roepstorff, 1974). All the VKD clotting factors require the 9–13 Glu residues at the

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Fig. 1 Vitamin K cycle. During vitamin K-dependent (VKD) carboxylation, the glutamate (Glu) of the VKD protein is converted to gamma-carboxyglutamate (Gla) by GGCX using vitamin K hydroquinone, carbon dioxide (CO2), and oxygen (O2) as cofactors. Concomitant with carboxylation, vitamin K hydroquinone is oxidized to vitamin K epoxide. Vitamin K epoxide is reduced to vitamin K by VKOR. The reduction of vitamin K to vitamin K hydroquinone is carried out by the as-yet-unidentified VKR.

N-terminus of the protein to be fully carboxylated in order for the protein to be functional. Defects of VKD carboxylation have long been known to cause bleeding disorders, known as combined vitamin K-dependent coagulation factors deficiency (VKCFD; Napolitano, Mariani, & Lapecorella, 2010; Prentice, 1985). With the discovery of new Gla proteins, the importance of VKD carboxylation has been extended beyond coagulation into a number of other physiological functions. For example, carboxylated matrix Gla protein (MGP) is a strong inhibitor of vascular calcification and of connective tissue mineralization, and uncarboxylated MGP has been implicated in cardiovascular diseases and other nonbleeding syndromes (Willems, Vermeer, Reutelingsperger, & Schurgers, 2014). Another VKD protein, osteocalcin (also called bone Gla protein, or BGP), is produced by osteoblasts and is important for bone formation (Ducy et al., 1996); recent studies suggest that osteocalcin also functions as a hormone affecting glucose metabolism (Mera et al., 2016). Therefore, functional study of the vitamin K cycle enzymes is not only essential for understanding blood coagulation, but it is also important for understanding many other physiological processes.

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1.1 Functional Study of VKOR by In Vitro Activity Assay The enzymatic activity of VKOR was discovered in 1970 (Bell & Matschiner, 1970). These authors showed that vitamin K-deficient rats can efficiently reduce KO to vitamin K and that this activity is sensitive to warfarin inhibition. Results from rat liver homogenates showed that the enzymatic activity of VKOR was enriched in purified microsomes (Zimmermann & Matschiner, 1974). However, fresh purified microsomes are inactive unless a thiol reagent, such as dithiothreitol (DTT), is included, suggesting that the thiol reagent is an artificial electron donor for VKOR activation. Based on a chemical model study, Silverman proposed that DTT reduces a critical disulfide bond within the active site of VKOR, and that the reduced cysteines are reoxidized to a disulfide during the reduction of KO to vitamin K (Silverman, 1981). Dithiol reagents were found to be more efficient than monothiol reagents as reductants for VKOR enzymatic activity (Lee & Fasco, 1984). The authors of this study suggested that monothiols are inefficient because a major proportion of the mixed disulfide, initially formed by reaction between the monothiol and the VKOR disulfide, undergoes reoxidation before a second molecule of monothiol can react. With the identification of the VKOR gene and functional studies of VKOR at the molecular level, we now know that Cys132 and Cys135 are the active site residues that form a disulfide after the reduction of KO (Li et al., 2004; Rost et al., 2004; Wajih, Hutson, Owen, & Wallin, 2005). The need to reduce an active site disulfide of VKOR to initiate its enzymatic activity stimulated the search for the physiological reductant of VKOR (Silverman & Nandi, 1988). Thijssen et al. reported that rat liver microsomes with VKOR also contain NADH-dependent lipoamide reductase activity (Thijssen, Janssen, & Vervoort, 1994). The reduced lipoamide stimulates microsomal VKOR activity with kinetics comparable to those of DTT. They proposed that microsomal lipoamide reductase is associated with VKOR in the membrane and is capable of transferring the reducing equivalents from NADH via reduced lipoamide to VKOR (Thijssen et al., 1994). However, a vast body of evidence suggests that thioredoxin is the physiological reductant of VKOR (Gardill & Suttie, 1990; Johan, van Haarlem, Soute, & Vermeer, 1987; Silverman & Nandi, 1988). Results from Soute et al. show that the protein disulfide isomerase (PDI) enhances the thioredoxin-driven VKOR activity
10-fold (Soute, Groenen-van Dooren, Holmgren, Lundstrom, & Vermeer, 1992). However, this hypothesis has been questioned by Preusch (1992), because thioredoxin is

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predominantly a soluble cytoplasmic protein that would not have access to the luminal surface of the ER where VKOR’s active site is located. Based on results showing that reduced RNAase can trigger PDI-dependent VKD carboxylation, and that this activity was inhibited by the PDI inhibitor bacitracin and by siRNA silencing of PDI, Wajih et al. proposed that PDI can directly provide electrons for the reduction of VKOR’s active site (Wajih, Hutson, & Wallin, 2007). Recently, by scanning a large number of mammalian thioredoxin-like ER proteins, including PDI, Schulman et al. reported that VKOR prefers the membrane-bound thioredoxin-like redox partners TMX (Schulman, Wang, Li, & Rapoport, 2010). Despite decades of effort, the physiological reductant of VKOR is still unknown. In the absence of a confirmed physiological reductant for VKOR, DTT is used as the reductant in the standard VKOR in vitro activity assay. This type of assay was first performed under a nitrogen atmosphere by blowing nitrogen over the incubation mixture (Whitlon, Sadowski, & Suttie, 1978; Zimmermann & Matschiner, 1974). The reaction was started by the addition of 3H-labeled KO and stopped by the addition of two volumes of an organic solvent, such as isopropanol:hexane (3:2). The radioactive K vitamins in the hexane extract were separated by reversed-phase silica-based thin-layer chromatography. KO and vitamin K spots were cut from the plate, placed in counting vials with scintillation fluid, and quantified by measuring the radioactivity with a liquid scintillation counter (Matschiner, Bell, Amelotti, & Knauer, 1970). Because of its complexity and its use of radioactive vitamin K, this type of VKOR activity assay was soon replaced; newer assays were performed using reverse-phase HPLC to separate the reactant and product of K vitamins, which were measured using UV detection (Elliott, Townsend, & Odam, 1980; Fasco & Principe, 1980). Typically, the reaction was performed with nonlabeled KO; after stopping the reaction with organic solvent, the vitamin K-containing organic phase was evaporated, redissolved, and applied to a reverse-phase HPLC using isocratic elution and UV detection at 250 nm. It has been suggested that the addition of silver nitrate to the extraction mixture prevents the extraction of DTT and thereby the formation of vitamin K-derived products on the HPLC column (Thijssen, 1987). To increase the accuracy of the assay, vitamin E acetate (Thijssen, Janssen, & DrittijReijnders, 1986) or vitamin K125 (Chu, Huang, Williams, & Stafford, 2006) was included in the extraction solvent as the internal standard. Over the past three decades, HPLC-based K-vitamin separation and detection has

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W/O DTT VKOR + DTT W/O VKOR

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KO

K

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1

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Fig. 2 Chromatogram of HPLC-based VKOR in vitro activity assay. The VKOR in vitro activity assay was performed using KO as the substrate and DTT as the artificial reducing agent. The amount of vitamin K (K) production was used to evaluate VKOR activity. K vitamins in the reaction mixture were extracted by organic solvent and separated by HPLC with UV detection at 250 nm. Negative controls were reactions that had no enzyme added (W/O VKOR) or that had no DTT added (W/O DTT). Vitamin E acetate was included in the organic extraction solvent as the internal standard (STD).

been the standard in vitro activity assay of VKOR for its functional study. Fig. 2 shows a typical chromatogram of the activity assay for VKOR’s reduction of KO to vitamin K. In addition to directly assaying VKOR’s ability to reduce KO or vitamin K, indirect approaches, coupling VKOR activity with VKD carboxylation, have also been used for functional studies of VKOR (refer to Section 1.3; Johan et al., 1987; Wallin & Martin, 1987). These in vitro studies have significantly contributed to our understanding of KO reduction, VKOR gene identification, VKOR active site identification, and recombinant VKOR purification and characterization. However, they have been unable to explain the correlation between VKOR mutations and their clinical phenotypes (Hodroge et al., 2012). Additionally, the use of DTT in these assays makes it impossible to explore the role of the physiological reductant for VKOR active site regeneration. To address these questions, an alternative activity assay to study VKOR function in the native milieu is required.

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1.2 Function Study of GGCX by In Vitro Activity Assay GGCX is a dual-function enzyme that accomplishes VKD carboxylation. During the process of carboxylation, the γ-hydrogen of the Glu residue of the substrate is abstracted, then CO2 is added. Simultaneously, GGCX oxidizes KH2 to KO to provide the energy required for the carboxylation (Larson, Friedman, & Suttie, 1981) (Fig. 1). The enzymatic activity of GGCX was first discovered in the 1970s (Girardot, Delaney, & Johnson, 1974; Shah & Suttie, 1974). It was found that radioactive 14CO2 was incorporated into prothrombin in rats, and that the amount of 14CO2 incorporation was increased three- to fourfold by the administration of vitamin K (Girardot et al., 1974). These authors concluded that VKD “completion” of prothrombin is actually carboxylation of Glu residues of the prothrombin precursor. This has been further confirmed by an in vitro assay using the postmitochondrial supernatant of rat hepatocytes (Shah & Suttie, 1974). Structural studies of bovine prothrombin have established that the 10 Gla residues are located at the protein’s N-terminal region (amino acid residues 1–42), now known as the Gla domain (Magnusson, Sottrup-Jensen, Petersen, Morris, & Dell, 1974). Stenflo et al. identified the structure of the Gla residues, and showed that Glu residues in prothrombin are modified by replacement of one hydrogen on the γ-carbon atom by a carboxyl group (Stenflo et al., 1974). These studies laid the foundation for the functional study of GGCX. The functional studies of GGCX were originally based on determining the incorporation of radioactive 14CO2 into the endogenous microsomal protein precursor substrates (Esmon, Sadowski, & Suttie, 1975). Detergentsolubilized rat liver microsome was shown to be an efficient system for GGCX function studies in vitro (Mack et al., 1976). As crude microsomal extracts from normal animals contain only a very small amount of endogenous protein substrates (mainly the precursors of prothrombin and factor X), treatment of animals with warfarin or feeding of animals on vitamin K-deficient diets was shown to be beneficial in obtaining substrate-enriched microsomal preparations for GGCX function studies (Vermeer, Soute, De Metz, & Hemker, 1982). Using this assay, it has been shown that the active form of vitamin K for GGCX is KH2, and that the enzymatic activity of GGCX is dependent on oxygen and bicarbonate (Sadowski, Esmon, & Suttie, 1976). Jones et al. reported that CO2 is the active specie participating in VKD carboxylation of Glu residues in the protein substrates (Jones, Gardner, Cooper, & Olson, 1977). VKD carboxylation has been associated

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with the oxidation of KH2 to KO (Sadowski, Schnoes, & Suttie, 1977), and a detailed stoichiometry study of Glu carboxylation and KO formation has confirmed that the two reactions are actually carried out by the same enzyme (Larson et al., 1981). Nowadays, the in vitro functional studies of GGCX were mainly carried out using artificial peptide substrates. One artificial substrate that has been used for GGCX function studies is the pentapeptide FLEEV (corresponding to residues 5–9), a peptide based on the bovine prothrombin protein sequence. It appears that the conditions for FLEEV carboxylation are identical with those for prothrombin carboxylation (Suttie & Hageman, 1976). By comparing the carboxylation of endogenous protein precursors and a variety of short peptide substrates, these authors showed that synthetic peptide substrates are carboxylated less effectively than endogenous protein substrates (Rich, Lehrman, Kawai, Goodman, & Suttie, 1981; Suttie, Lehrman, Geweke, Hageman, & Rich, 1979). In addition, a fivefold higher vitamin K concentration is required for half-maximal carboxylation of the peptide substrates than the protein substrates. GGCX carboxylates only glutamic acid residues, not aspartic acid and other related residues. It appears that hydrophobic residue around sequence EE is associated with enhanced substrate activity. As the addition of detergent to intact microsomes resulted in a 10- to 20-fold stimulation in carboxylation of a peptide substrate, it was proposed that the active site of the GGCX is accessible only from the lumen of the microsomal membrane (Carlisle & Suttie, 1980). The most popular substrate used for GGCX functional study in vitro is FLEEL (corresponding to residues 5–9 of rat prothrombin protein sequence). However, FLEEL is a poor substrate analog for GGCX for the following reasons: (1) of the two glutamic acids, only the first glutamic acid of FLEEL is carboxylated during in vitro activity assay (Burgess, Esnouf, Rose, & Offord, 1983; Decottignies-Le Marechal, Rikong-Aide, Azerad, & Gaudry, 1979); (2) only small amounts of substrate are converted to product (Soute, Ulrich, & Vermeer, 1987); and (3) FLEEL binds poorly to GGCX, with a Km of
4 mM (Suttie et al., 1979). Knowing that GGCX interacts with clotting-factor precursors, mainly through an 18-amino acid propeptide (Jorgensen et al., 1987; Pan & Price, 1985), and since propeptide significantly increases small peptide substrate carboxylation (Knobloch & Suttie, 1987), Ulrich et al. used a 28-residue peptide, based on residues 18 to +10 in prothrombin (proPT28), as the peptide substrate of GGCX (Ulrich, Furie, Jacobs, Vermeer, & Furie, 1988). This 28-residue peptide is efficiently carboxylated, with a Km that is three orders of magnitude lower

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than FLEELs. In addition to the 28-residue peptide, a 59-residue peptide, containing the propeptide sequence and the whole Gla domain (residues 18 to 41) of factor IX (proFIX59), was also proved to be an efficient substrate for in vitro carboxylation study; the apparent Km (0.55 μM) of proFIX59 is
five times lower than that of proPT28 (Wu, Soute, Vermeer, & Stafford, 1990). Finally, decarboxylated BGP and MGP are also good substrates for GGCX function studies (Engelke, Hale, Suttie, & Price, 1991; Vermeer, Soute, Hendrix, & de Boer-van den Berg, 1984). GGCX activity assays are mainly based on the incorporation of 14CO2 into VKD protein precursors or synthetic peptides. The 14CO2 incorporation into the γ-[14C]carboxyglutamate product is then determined in either the trichloroacetic acid (TCA) insoluble fractions (for the protein substrates) or the TCA soluble fractions (for the peptide substrates). After terminating the reaction with TCA, investigators must remove the unreacted 14CO2 by manipulating the assay tubes containing volatile 14CO2 in potentially unsafe circumstances: centrifugation (Esmon et al., 1975) or pipetting and vigorous bubbling of the samples with carbon dioxide (Suttie & Hageman, 1976). In order to optimize safety, accuracy, and simplicity, the method for determining 14CO2 incorporation into the peptide substrates was modified by instead boiling the TCA-treated samples in an exhaust hood (Romiti & Kappel, 1985; Ulrich et al., 1988). Recently, Kaesler et al. developed a nonradioactive assay for GGCX function study that used a fluorescein isothiocyanate (FITC)-labeled hexapeptide (FLEELK-FITC) that can be readily separated and detected in its unmodified and γ-glutamyl carboxylated form by reversed-phase HPLC with fluorescence detection (Kaesler et al., 2012). In addition to the enzymatic catalysis assays used for GGCX function studies, Presnell et al. described a fluorescence assay for studying the interaction between GGCX and propeptide (Presnell, Tripathy, Lentz, Jin, & Stafford, 2001). This assay allows measurement of the number of propeptide-binding sites, the equilibrium dissociation constant for the propeptide–GGCX interaction, and the off-rate of a propeptide from the enzyme. These authors show that the off-rate for the FIX propeptide is 3000-fold slower than the rate of carboxylation, a difference that may explain how GGCX can carry out multiple carboxylations of a substrate during the same binding event. In order to stabilize GGCX and provide a native-like environment for its function study, GGCX was recently incorporated as single molecules into nanodiscs (Fig. 3). The assembled GGCX nanodiscs are structurally stable and catalytically active (Hebling et al., 2010). Hydrogen/deuterium exchange mass spectrometry was used to characterize

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Fig. 3 Schematic diagram of GGCX incorporated into nanodisc. GGCX, a fivetransmembrane domain integral membrane protein, was incorporated into lipid bilayers with the help of membrane scaffold protein (MSP) to form the nanodisc which renders GGCX soluble in aqueous solution and structurally stable.

specific regions of nanodisc-incorporated GGCX that exhibited structural rearrangements upon binding to propeptides. Results from these studies showed that the propeptide-binding region is located in the C-terminal of GGCX (Hebling et al., 2010; Parker et al., 2014); this is consistent with regions of propeptide binding previously identified by peptide-based affinity labeling and enzyme kinetic studies. Most of our knowledge about GGCX’s function has been obtained from these in vitro studies carried out under artificial conditions. Thus, we lack understanding of how GGCX carboxylates the natural VKD protein substrates in the native milieu. For example, it is not clear why some of the GGCX mutations found in patients cause carboxylation deficiencies in coagulation factors (which result in bleeding disorders), while others cause undercarboxylation of extrahepatic VKD proteins such as MGP (which results in vascular calcification). Therefore, it is essential to develop a better activity assay to explore GGCX function in its native environment in order to clarify the complex mechanism of this multisubstrate enzyme.

1.3 Function Study of VKR by In Vitro Activity Assay The enzyme that reduces vitamin K to KH2 for VKD carboxylation is VKR. It has been proposed that in vitro, vitamin K can be reduced to KH2 via two pathways: a DTT-supported warfarin-sensitive pathway, accomplished by

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VKOR, and an NAD(P)H-dependent warfarin-resistant pathway, catalyzed by NAD(P)H quinone oxidoreductase 1 (NQO1) (Wallin & Martin, 1987; Whitlon et al., 1978). However, there is no in vivo experimental evidence to support this dual-pathway hypothesis. When VKOR is inactivated by warfarin or knocked out by gene targeting in HEK293 cells, those cells can still efficiently reduce vitamin K to support VKD carboxylation (Tie, Jin, Straight, & Stafford, 2011; Tie, Jin, Tie, & Stafford, 2013). Additionally, a phenotype of VKOR-deficient mice that died due to extensive intracerebral hemorrhage could be rescued by the oral administration of vitamin K (Spohn et al., 2009). These in vivo studies suggest that VKOR is not the key player for vitamin K reduction in physiological conditions. NQO1 is a ubiquitous dicoumarol-sensitive flavoprotein that can catalyze the two-electron reduction of several quinones (including vitamin K) to hydroquinones (Fasco & Principe, 1982). It has long been proposed as the warfarin-resistant enzyme for vitamin K reduction (Ernster, Lind, & Rase, 1972); however, NQO1-knockout mice do not have bleeding problems (Gong, Gutala, & Jaiswal, 2008), and NQO1-deficient mice survived at the same frequency as wild-type mice when poisoned with warfarin (Ingram, Turbyfill, Bledsoe, Jaiswal, & Stafford, 2013); these in vivo results suggest that NQO1 does not play a major role in the production of KH2 to support VKD carboxylation. Despite decades of research into VKR, its identity is still unknown. However, there are several lines of evidence that demonstrate the existence and the importance of VKR. Most notably, patients that are overdosed with warfarin can be rescued by large doses of vitamin K through the action of a warfarin-resistant enzyme, designated the “antidotal enzyme,” that reduces vitamin K. The antidotal effect of vitamin K was first discovered in 1966 (O’Reilly & Aggeler, 1966) and was recently demonstrated in the VKORknockout mouse (Spohn et al., 2009) and in a cell-based vitamin K cycle enzyme study (Tie et al., 2011). In addition, we have observed, in AV12 cells derived from Syrian golden hamsters, a warfarin-sensitive VKR activity that is distinct from VKOR (Tie et al., 2011). A detailed understanding of the characteristics of both the warfarin-resistant and warfarin-sensitive VKRs will not only help us to understand the mechanism of vitamin K reduction, but will also be important for regulating blood coagulation and other mechanisms of physiological homeostasis. As in the study of VKOR, function studies of VKR can be performed by directly measuring the reduction of vitamin K to KH2 using a HLPC-based assay (Fasco & Principe, 1980). As KH2 is unstable and can easily be oxidized

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back to vitamin K during extraction and isolation, KH2 was stabilized in one of two ways: by including butylated hydroxytoluene in the vitamin K extraction buffer (Thijssen, Soute, Vervoort, & Claessens, 2004) or by performing the reaction and subsequent vitamin K isolation in sealed vials under an N2 environment (Rishavy et al., 2013). The ability of purified recombinant VKOR to reduce vitamin K to KH2 has been studied by directly measuring KH2 production (Chu et al., 2006). Results show that VKOR can reduce vitamin K using the same active site cysteine residues that it uses to reduce KO, and that both activities are sensitive to warfarin inhibition (Jin, Tie, & Stafford, 2007). Rishavy et al. reported that VKOR exists as a dimer, with one molecule that reduces KO and another that reduces vitamin K (Rishavy et al., 2013). Furthermore, it has been shown that VKOR-like enzyme (VKORL, a paralog enzyme of VKOR) can also reduce vitamin K to KH2 (Westhofen et al., 2011). Despite the fact that VKOR can function as VKR in vitro, it appears that in physiological conditions VKOR is not the main contributor for vitamin K reduction (Tie et al., 2011). In addition to directly measuring KH2 production, VKR function studies have also been carried out using an indirect method that coupled vitamin K reduction with VKD carboxylation and determined the VKRsupported incorporation of 14CO2 either into the endogenous VKD protein substrate (Wallin, Gebhardt, & Prydz, 1978) or into the synthetic peptide substrates (Johan et al., 1987; Wallin & Martin, 1987). Microsomes prepared from animal liver were used to mimic the in vivo vitamin K metabolism in vitro. Using this coupled in vitro activity assay system, Wallin et al. have confirmed the existence of both the warfarin-sensitive and warfarin-resistant pathways for vitamin K reduction (Wallin & Martin, 1987). In addition, these authors have shown that the reduction of vitamin K is the rate-limiting step in VKD carboxylation (Wallin, Sane, & Hutson, 2002), which has been further confirmed by studying VKD coagulation factor carboxylation in mammalian cells (Hallgren, Qian, Yakubenko, Runge, & Berkner, 2006; Sun, Jin, Camire, & Stafford, 2005; Wajih et al., 2005). Recently, Rishavy et al. reported a similar assay system that coexpressed VKOR and GGCX in insect cells lacking endogenous VKD carboxylation components (Rishavy et al., 2013). VKR function was studied using the coupled activity assay system from intact microsomes. This in vitro coupling-assay approach appears reasonable for studying vitamin K metabolism and VKD carboxylation; however, the enzymes of the vitamin K cycle have very different lipid and/or detergent requirements for their activities (Carlisle & Suttie, 1980;

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Hildebrandt, Preusch, Patterson, & Suttie, 1984), which makes it difficult to study the interactions and activities of these enzymes together in vitro. As for VKOR and GGCX, an activity assay for VKR in physiological conditions would help to understand the role of this enzyme in the complex vitamin K cycle. However, because VKR’s identity is unknown, establishing such an assay is even more challenging. We hope that the recently emerged genome-scale CRISPR-Cas9 knockout screening will accelerate the identification of VKR. Detailed insights into the structure–function relationships of VKR will not only extend our understanding of the mechanism of vitamin K reduction, but will also help us to develop new ways of regulating coagulation and treating thrombosis.

2. CELL-BASED ASSAY FOR FUNCTIONAL STUDY OF VITAMIN K CYCLE ENZYMES 2.1 Rationale To better understand VKD carboxylation in the native milieu, we developed a cell-based reporter assay system for functional study of the vitamin K cycle enzymes (Tie et al., 2011). In this system, we expressed a chimeric VKD reporter protein in mammalian cells; its carboxylation status is easily measured by enzyme-linked immunosorbent assay (ELISA). The efficiency of carboxylation of the reporter protein was used to determine the functionality of vitamin K cycle enzymes. This approach has the advantage of assessing the functionality of the vitamin K cycle enzymes in an environment that requires the enzymes to interact with their physiologic partners and/or other components of the vitamin K cycle. We used two mammalian cell lines, HEK293 and AV12, which have all the necessary components for VKD carboxylation. The HEK293 cells can efficiently carboxylate VKD proteins and have been commonly used to produce commercial therapeutic recombinant coagulation factors (Dumont, Euwart, Mei, Estes, & Kshirsagar, 2016). The AV12 cells, on the other hand, are less efficient in the carboxylation of VKD proteins (Yan et al., 1990), which could suggest that these cells use different vitamin K pathways for making VKD proteins. By expressing an appropriate VKD reporter protein in these two cell lines, and by manipulating factors that affect VKD carboxylation, we hoped to determine the contributions of the various enzymes in vitamin K cycle. In the HEK293 and AV12 cells, we stably expressed a chimeric VKD reporter protein, protein C with its Gla domain exchanged with that of factor IX (FIXgla-PC). The functionality of the vitamin K cycle enzymes is

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evaluated by culturing the reporter cells under various conditions and by using ELISA to measure the amount of carboxylated reporter protein secreted in the cell culture medium (Fig. 4). The rationale for using FIXglaPC as the reporter protein is that the uncarboxylated protein C undergoes selective intracellular degradation through a quality control mechanism in the ER (Tokunaga, Takeuchi, Omura, Arvan, & Koide, 2000; Tokunaga, Wakabayashi, & Koide, 1995) that decreases the background secretion of reporter protein; FIXgla enables the ready quantitative detection of the carboxylated reporter protein by a calcium-dependent conformation-specific monoclonal antibody (α-FIXgla mAb) that recognizes only the fully carboxylated FIXgla (Feuerstein et al., 1999). Additionally, using the sandwich-based ELISA (Fig. 4C) to detect the carboxylated chimeric reporter protein eliminates the interference of the endogenous protein C or FIX in the cell culture medium. Furthermore, ELISA’s detection of the reporter protein can be easily adapted with high-throughput screening of potential inhibitors of the vitamin K cycle enzymes. Recently, similar cell-based assays have been performed using different reporter proteins (Fregin et al., 2013; Haque, McDonald, Kulman, &

Fig. 4 Cell-based activity assay for vitamin K cycle enzymes with ELISA detection. (A) Domain structure of reporter protein FIXgla-PC. AP, activation peptide of protein C; Catalytic domain, region containing the serine protease catalytic triad of protein C; EGF, region of protein C homologous to human epidermal growth factor; FIXgla, Gla domain of human factor IX. (B) Schematic diagram of FIXgla-PC/HEK293 reporter cell. Reporter protein FIXgla-PC is carboxylated by the endogenous vitamin K cycle enzymes of the HEK293 cell in the presence of vitamin K. The carboxylated FIXgla-PC is secreted from HEK293 cell into the cell culture medium. (C) Sandwich-based ELISA detection of the carboxylated reporter protein. Carboxylated FIXgla-PC is captured by α-FIXgla mAb (capture antibody) in the presence of Ca2+ and detected by HRP-conjugated sheep antihuman protein C antibody (detector antibody).

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Rettie, 2014). Haque et al. engineered an HEK293-derived reporter cell line (HEK293-C3) that expresses a chimeric reporter protein, F9CH, which is a VKD integral membrane protein (human proline-rich Gla protein 2) with its Gla domain replaced by FIXgla (Haque et al., 2014). This chimeric reporter protein is present mainly on the cell surface, with its FIXgla domain exposed outside the cell. Carboxylated reporter proteins were detected by staining the cells with fluorescein-labeled α-FIXgla mAb, which only recognizes carboxylated FIXgla and can be quantified using flow cytometric detection. Instead of using chimeric proteins as reporters, Fregin et al. used full-length FIX as the reporter protein in HEK293T cells; they used the activity of secreted FIX in the cell culture medium to evaluate the efficiency of VKD carboxylation (Fregin et al., 2013). It should be noted that, unlike the other two assays, Fregin’s assay requires transient expression of the reporter protein in each individual sample, which can create larger variations in reporter protein expression between samples. Unlike the traditional in vitro activity assays, these cell-based assays provide a platform for the functional study of enzymes in the vitamin K cycle within a cellular milieu, where all the endogenous components required for VKD carboxylation are present. Therefore, results obtained from these assays for vitamin K cycle enzymes are expected to reflect their functions in physiological conditions.

2.2 Establishment of the FIXgla-PC Reporter Cell Lines To construct the FIXgla-PC chimeric reporter protein, the cDNA encoding human protein C was amplified by PCR using pCMV-SPORT6-protein C (Open Biosystems, Huntsville, AL) as a template. A BstBI (TTCGAA) restriction site was introduced at the end of protein C’s propeptide (amino acid residues -4I and -3R) using codon degeneracy. The Gla domain of protein C (residues 1–46) was removed by restriction digestion using BstBI and BstEII (a unique restriction site at the end of protein C’s Gla domain) and replaced with the PCR fragment encoding FIX’s Gla domain flanked by the same restriction sites. This chimeric FIXgla-PC fusion was subcloned into the mammalian expression vector pcDNA3.1 (Invitrogen, Carlsbad, CA) using the XbaI site to generate the reporter protein expression vector pcDNA3.1-FIXgla-PC. The FIXgla-PC reporter protein was stably expressed in HEK293 or AV12 cells. Cells were transfected with pcDNA3.1-FIXgla-PC plasmid DNA using Lipofectamine (Invitrogen, Carlsbad, CA) according to the

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manufacturer’s protocol. Twenty-four hours posttransfection, cells were detached and diluted at 1:10 with fresh complete growth medium (DMEM/F-12 supplemented with 10% FBS and 1 Penicillin/Streptomycin) and plated into a 10-cm dish. The following day, selective medium with 300 μg/mL of hygromycin was added and cells were cultured at 37°C in the CO2 incubator until cells that did not have the plasmid DNA incorporated into their genomic DNA were killed by hygromycin. Surviving colonies were manually picked and amplified in a 24-well plate in order to screen for stable expression of the reporter protein. When wells were
70% confluent, 11 μM vitamin K was included in the complete growth medium and incubated for 48 h. Cell culture medium was collected and used directly for quantification of the secreted carboxylated FIXgla-PC by ELISA. The colony with the highest FIXgla-PC production was selected as the stable cell line to be used for reporter protein expression. To further purify the manually picked colonies, the colony with the highest FIXgla-PC expression was further screened using ClonePix FL technology (CDI Bioscience, Madison, WI). Cells were partially immobilized by plating in the semisolid medium to form single colonies. After treatment with vitamin K, carboxylated reporter protein FIXgla-PC that was secreted from the colonies, which was trapped in the vicinity of the colonies by the semisolid medium, was probed by fluorescein-labeled α-FIXgla mAb. Colonies that highly express FIXgla-PC were isolated using ClonePix FL, expanded, and characterized by ELISA. Characterization of the selected colonies was performed by culturing the cells under different conditions. As expected, secretion of carboxylated reporter protein increased with the increasing concentrations of vitamin K in the cell culture medium. When these cells were fed with KO, carboxylation of the reporter protein was significantly inhibited by warfarin, a VKOR inhibitor (Fig. 5).

2.3 ELISA-Based Evaluation of the Reporter Protein Carboxylation In this cell-based assay, the functions of vitamin K cycle enzymes were evaluated by determining the carboxylated FIXgla-PC in the cell culture medium directly, using ELISA (Tie et al., 2011). Reporter cells were seeded at
60% confluence in a 24-well plate. The following day, the cell culture medium was changed to complete growth medium, which was supplemented with either the desired substrates (such as KO or vitamin K) or with an inhibitor (such as warfarin). The cell culture medium was

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Fig. 5 Characterization of FIXgla-PC/HEK293 reporter cells with vitamin K and warfarin. FIXgla-PC/HEK293 reporter cells were cultured for 48 h with increasing concentrations of vitamin K (•) or 5 μM KO with increasing concentrations of warfarin (♦). The concentration of carboxylated FIXgla-PC in the cell culture medium was measured by ELISA.

collected and used directly for quantification of the secreted carboxylated FIXgla-PC by ELISA after 48 h incubation. ELISA plates were coated overnight at 4°C with 100 μL/well of α-FIXgla mAb (GMA-001, Green Mountain Antibodies, Burlington, VT). The concentration of the coating antibody was 2 μg/mL in 50 mM carbonate buffer (pH 9.6). After being washed five times with TBS-T washing buffer (20 mM Tris–HCl, pH 7.6, 150 mM NaCl, and 0.1% Tween 20), the plate was blocked with 0.2% bovine serum albumin in TBS-T washing buffer for 2 h at room temperature. Samples and protein standards (0.12–250 ng/mL) containing 3 mM CaCl2 were added at 100 μL/well and incubated for 2 h at room temperature. After being washed with TBS-T washing buffer containing 3 mM CaCl2, 100 μL of 1:2500 diluted sheep antihuman protein C IgG conjugated to horseradish peroxidase (SAPC-HRP, Affinity Biologicals Inc., Ancaster, ON, Canada) was added to each well and incubated for 45 min at room temperature. As α-FIXgla mAb is calcium dependent, it was important to include 3 mM Ca2+ in any part of the procedure in which the antibody and the antigen were bounded. After the unbound detecting antibody was washed off, 100 μL of 2,20 -azino-bis(3-ethylbenzthiazoline-6-sulfonic acid; ABTS) solution

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(KPL, Gaithersburg, MD) was added to each well and the absorbance was determined at 405 nm with a ThermoMax microplate reader (Molecular Devices, Sunnyvale, CA). To obtain the protein standard for ELISA, carboxylated FIXgla-PC chimeric protein was purified by affinity chromatography. Anticarboxylated FIXgla mAb was coupled to Affi-Gel 10 (Bio-Rad Laboratories Inc., Hercules, CA) according to the manufacturer’s protocol. HEK293 cells stably expressing a high level of FIXgla-PC reporter protein were cultured in complete growth medium supplemented with 11 μM vitamin K. The medium was collected after 48 h of incubation. Calcium chloride is added to the collected medium, to a final concentration of 3 mM. Five hundred milliliters of the collected medium were incubated with 1.5 mL of the prepared anticarboxylated FIXgla affinity resin overnight at 4°C with gentle rotation. The beads were spun down and packed into a 1.5  10 cm column. The column was washed first with buffer containing 20 mM Tris–HCl, 500 mM NaCl, and 3 mM CaCl2, and then with 20 mM Tris–HCl, 100 mM NaCl, and 3 mM CaCl2. Carboxylated FIXgla-PC reporter protein was eluted with 20 mM Tris–HCl, 100 mM NaCl, and 10 mM EDTA. The concentration of the eluted protein was calculated from its absorbance at 280 nm according to the extinction coefficient of 1%E280nm ¼ 14.5 (Kisiel & Davie, 1981). With the affinity-purified carboxylated FIXgla-PC as the standard protein, the standard curve for determining the carboxylated reporter protein with ELISA was plotted using a logit–log curve-fitting model (Plikaytis, Turner, Gheesling, & Carlone, 1991). In this model, logit OD405 is plotted against the log standard protein-dilution scale. Where p represents a proportion, the logit of that proportion can be obtained using the equation logit ( p) ¼ log [p/(1  p)]. Therefore, logit OD405 ¼ log [OD405/ (1  OD405)]. Fig. 6 shows a typical standard curve of a serial dilution of purified carboxylated FIXgla-PC, plotted by the logit-log curve-fitting model. The linear range for detection of the carboxylated reporter protein is between 0.24 and 125 ng/mL (γ ¼ 0.9991).

2.4 Applications The established cell-based assay allows one to study the vitamin K cycle enzymes in their native milieu. Two reporter cell lines can be used depending on the question being addressed. The HEK293 reporter cells have endogenous KO reductase activity that is warfarin sensitive and VKR

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0.5

–1

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–0.5 y = 0.9047x – 1.6973 R2 = 0.9991

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Fig. 6 Standard curve for measuring carboxylated reporter protein FIXgla-PC by ELISA. Affinity-purified carboxylated FIXgla-PC was serially diluted and determined by ELISA. Protein concentration and absorption at 405 nm were plotted using a logit–log curvefitting model.

activity that is warfarin resistant; AV12 reporter cells do not have warfarinresistant VKR activity. By supplementing with either KO or vitamin K as the substrate, these cell lines can be used to study the function of enzymes involved in the redox cycling of vitamin K. To study the two-step reduction of KO to KH2 by the enzymes in the vitamin K cycle (Fig. 1), FIXgla-PC/HEK293 reporter cells were cultured with complete growth medium containing either KO or vitamin K. Similar levels of carboxylated reporter protein were secreted independently of the substrate (vitamin K or KO) fed to the cells (Tie et al., 2011), indicating that these cells can efficiently reduce KO to vitamin K and vitamin K to KH2 to support VKD carboxylation. This allows one to differentiate the functions of the various enzymes involved in two-step reduction of vitamin K in the cycle. With KO as the substrate, FIXgla-PC carboxylation was completely inhibited by 2 μM warfarin, which suggests that VKOR, the molecular target for warfarin, is responsible for the reduction of KO in vivo. On the other hand, when vitamin K was the substrate, the reporter cells produced unaffected levels of carboxylated reporter protein in the presence of warfarin, suggesting that with enough vitamin K in the medium, inactivation of VKOR by warfarin does not affect the conversion of vitamin K to KH2. This result suggests that there must be a warfarin-resistant enzyme that

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reduces vitamin K to KH2 in HEK293 cells, data that agree with the antidotal effect of high-dose supplementation with vitamin K upon patients poisoned with warfarin. This cell-based assay has also clarified that the warfarinresistant pathway for vitamin K reduction in HEK293 cells is not accomplished by NQO1, the previously proposed antidotal enzyme. In AV12 cells, however, warfarin significantly inhibited FIXgla-PC carboxylation when vitamin K was used as the substrate (Tie et al., 2011). This suggests that vitamin K reduction in AV12 cells is carried out mainly via a warfarinsensitive pathway. It has been shown that, in vitro, VKOR can also reduce vitamin K through warfarin-sensitive pathways (Jin et al., 2007). To test the ability of VKOR to reduce vitamin K to KH2 in vivo, tyrosine 139 in VKOR was mutated to phenylalanine (Y139F), which converts VKOR to a warfarin-resistant form (VKOR-Y139F). This permitted the inactivation of the endogenous VKOR by warfarin while testing the function of the exogenously expressed VKORY139F. Because AV12 cells do not have warfarin-resistant enzyme(s) for vitamin K reduction, this cell line is a useful tool for studying vitamin K reduction by VKOR-Y139F. AV12 reporter cells expressing VKORY139F failed to produce carboxylated FIXgla-PC when fed KO in the presence of warfarin, suggesting that VKOR has very limited ability to convert vitamin K to KH2 in vivo (Tie et al., 2011). The strategy of using VKOR-Y139F to study VKOR function in the cell-based assay allowed us to clarify the disputed function of the conserved loop cysteines in human VKOR and to explore the function of bacterial VKOR homologs (VKORHs). It has been proposed that, like VKORHs, the two conserved loop cysteines of human VKOR shuttle electrons to its active site cysteines through an intramolecular electron transfer pathway (Li et al., 2010; Schulman et al., 2010). To test this hypothesis using the established cell-based assay, we first determined whether VKORHs could function in the mammalian vitamin K cycle to support VKD carboxylation. We expressed five VKORHs in HEK293 and AV12 reporter cell lines. Results show that Mt-VKORH (a VKORH from Mycobacterium tuberculosis) can efficiently support reporter-protein carboxylation using either KO or vitamin K as a substrate. Mutating both loop cysteines to alanine in MtVKORH or in human VKOR had only a minor effect on the activity of reporter-protein carboxylation, suggesting that these loop cysteines are not required for vitamin K cycling (Tie, Jin, & Stafford, 2012b). However, the loop cysteines of Mt-VKORH are essential for its activity in disulfide bond formation during protein folding in Escherichia coli (Wang, Dutton,

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Beckwith, & Boyd, 2011). These results together suggest that, in their respective native environments, human VKOR and VKORHs employ different mechanisms for active site regeneration. However, when MtVKORH is in the mammalian cell system, it employs a mechanism similar to that of human VKOR. Using the reporter cell line HEK293-C3, Haque et al. have shown that S-warfarin (Fig. 7) is
10 times more potent than R-warfarin for inhibiting VKD carboxylation (Haque et al., 2014), which is consistent with the potencies of warfarin enantiomers as oral anticoagulant drugs. In addition, these authors have studied the structure–activity relationship of warfarin metabolites used for inhibition of VKD carboxylation. Results from this cell-based assay, which agree with the early in vivo studies, show that all of the monohydroxy metabolites of warfarin (except for 10-hydroxywarfarin and warfarin alcohols) show less potency for inhibiting VKD carboxylation. Based on their results, these authors proposed that steric constraints for binding of warfarin to its target protein VKOR are greatest around the C-7 and C-8 positions, but that much more flexibility exists in the region of the VKOR active site that contacts the C-6 position (Fig. 7). These cell-based assays allowed us to access the physiological functions of the vitamin K cycle enzymes in a way not previously possible. The strategy of creating a warfarin-resistant version of VKOR makes it possible to explore VKOR’s function in the cellular environment by inactivating the endogenous VKOR with warfarin (Tie, Jin, & Stafford, 2012a; Tie et al., 2012b). However, the disadvantage of this strategy is that all the experiments must be performed in the presence of warfarin. This makes it impossible to study the warfarin resistance of the naturally occurring VKOR mutations, and it potentially compromises functional study of the wild-type enzyme. Therefore, functional study of these enzymes with a clean background requires knockout of the endogenous enzymes in the reporter cells.

Fig. 7 Chemical structure of warfarin enantiomers.

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3. FUNCTIONAL STUDY OF VKOR USING TALENsMEDIATED GENE KNOCKOUT REPORTER CELLS 3.1 Rationale VKOR is the enzyme that reduces KO in the vitamin K cycle (Fig. 1). It is the target of warfarin, the most widely used oral anticoagulant for prevention and treatment of thrombosis. However, achieving the desired anticoagulation is challenging because of warfarin’s narrow therapeutic index and the broad individual variability of dosing requirements. With the discovery of the gene that encoding VKOR (Li et al., 2004; Rost et al., 2004), studies suggest that genetic variations in VKOR significantly contribute to warfarin dosage variability (Lee & Klein, 2013; Zineh, Pacanowski, & Woodcock, 2013). It is estimated that
30% of patients receiving warfarin would benefit from VKOR pharmacogenetics modeling at the beginning of warfarin therapy (Eriksson & Wadelius, 2012). VKOR pharmacogenetics is thought to be clinically useful enough that, in 2010, the US Food and Drug Administration revised the warfarin product labels to include VKOR genotypes in the warfarin dose recommendations (Administration & U. S. F. a. D., 2010). Missense mutations that have been identified in the VKOR coding regions of warfarin-resistant patients are believed to produce VKOR molecules that are more resistant to warfarin inhibition, thus requiring higher therapeutic warfarin doses. The traditional DTT-driven VKOR activity assay has been used to evaluate the effect of these mutations on VKOR activity and its warfarin resistance. However, except for the mutation at residue 139 in human VKOR, the results from in vitro studies of other VKOR mutations cannot explain the warfarin-resistant phenotype (Pelz et al., 2005; Rost et al., 2004). Recently, Hodroge et al. systematically studied all the naturally occurring VKOR mutants identified in patients requiring high dosages of oral anticoagulants (Hodroge et al., 2012). They expressed these VKOR mutants in Pichia pastoris and prepared VKOR-containing microsomes for functional studies using an in vitro DTT-driven activity assay. This study showed that of the 25 VKOR mutants, only 6 had increased resistance toward anticoagulant drugs; 10 had essentially no in vitro activity, which makes it difficult to explain how these VKOR variants function in vivo. These discrepancies presumably arise from the in vitro activity assay itself, which was performed in artificial conditions using the nonphysiological reductant DTT.

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The cell-based assay in Section 2 has the advantage of studying the function of VKOR in its native milieu with all the required endogenous components. The disadvantage, however, is that the endogenous enzymes provide a high background. Therefore, it is not appropriate to directly use this cell-based assay to evaluate the functional consequences of VKOR mutations. To overcome this problem, the endogenous VKOR gene was knocked out in the reporter cells using transcription activator-like effector nucleases (TALENs)-mediated genome editing (Sanjana et al., 2012). Targeted gene knockout is achieved through the DNA repair machinery’s action upon the chromosomal DNA double-strand breaks (DSBs) caused by TALENs in the reporter cells (Fig. 8A). TALENs consist of a pair of DNA cleavage domains of endonuclease FokI that have been fused to customized DNA-binding domains that recognize a specific region of the targeted gene sequence. When TALENs bind to the target sequences, the fused FokI are brought close enough to form a homodimer in the target spacer region. This homodimer of FokI is the active form, which will create DSBs in the genomic DNA. The repair of the DSBs in mammalian cells by nonhomologous

Fig. 8 Schematic diagram of TALENs-mediated gene knockout. (A) Customized DNAbinding domain of TALE recognizing a specific genomic DNA sequence is fused to the catalytic domain of FokI endonuclease (TALEN). When a pair of TALENs recognize the top and bottom strands of the target sites flanking a 14–20 bp spacer, the FokI dimerizes and cuts the DNA in the spacer region to create the double-strand breaks (DSBs). (B) When DSBs in a gene locus are repaired by the error-prone nonhomologous end-joining (NHEJ) pathway, it leads to the introduction of variable length insertion or deletion (indel) mutations resulting in ablation of the targeted gene function.

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end-joining (NHEJ) pathways introduces mutations via random insertions or deletions (indel mutations; Fig. 8B). This often produces frameshift mutations that ablate the targeted gene function. Based on loss-of-function screening and characterization of TALENsmediated gene targeting in the above reporter cells, both VKOR and VKORL must be knocked out to eliminate the ability of HEK293 cells using KO as substrate to support VKD carboxylation (Tie et al., 2013). Reporter cells with both VKOR and VKORL knocked out (double-gene knockout, or DGKO) provide an ideal tool for VKOR function study with a clean background.

3.2 Establishment of the TALENs-Mediated VKOR/VKORL Knockout Reporter Cell Line To knock out the endogenous VKOR and VKORL genes in HEK293 cells, TALENs recognition sites that target exon 1 (near the start codon region of human VKOR and VKORL) in the genomic DNA sequences were designed using TALE-NT 2.0 software (Doyle et al., 2012). Both the left and the right TALENs were designed to target a 20-bp sequence, with 17-bp spacing for VKOR and 15-bp spacing for VKORL (Fig. 9). The 18-mer DNA-binding arrays of TALEN were engineered using TALE Toolbox, as described previously (Sanjana et al., 2012), with minor modification. Instead of generating 72 monomers (18 monomers for each TALE DNA-binding domain), 40 monomers were created using PCR to avoid monomer repeats. PCR products for making TALENs DNA-binding arrays were purified using gel purification. The assembled TALENs were cloned into the corresponding pTALEN_v2 vectors provided in the TALE Toolbox kit. The integrity of each assembled TALEN was verified by sequence analysis.

Fig. 9 TALENs target sequences for VKOR and VKORL knockout. The left and right TALENs binding sequences for VKOR (top) and VKORL (bottom) are highlighted. The start codon for protein translation is indicated by arrows.

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The reporter cell line of FIXgla-PC/HEK293 was used for TALENsmediated VKOR gene knockout. Plasmid DNA containing the left and right TALENs targeting VKOR was transiently expressed in FIXgla-PC/ HEK293 cells. Forty-eight hours posttransfection, cells were treated with trypsin, filtered with a 30-μm filter (Sysmex Partec GmbH, Gorlitz, Germany), and diluted with complete growth medium to obtain single-cell suspensions. Diluted cells were seeded into 96-well plates for functional screening of the gene knockout. Alternatively, single-cell suspensions of the transfected cells were plated into 150-mm dishes. Single colonies were manually picked for functional screening. VKOR is the enzyme responsible for reducing KO to vitamin K, while HEK293 cells possess an “antidotal” enzyme that can reduce vitamin K to KH2. We reasoned that HEK293 cells with the VKOR gene knockout would abolish only the KO reductase activity but leave the K reductase activity intact (Fig. 1). Therefore, functional screening of VKOR knockout cell colonies was performed by first measuring the efficiency of reporterprotein carboxylation when the cell colonies were fed with 5 μM KO for 48 h. Carboxylated FIXgla-PC in the cell culture medium was measured by ELISA, as described in Section 2.3. Cell colonies with significantly reduced carboxylation of reporter protein were then fed with 11 μM vitamin K for 24 h to test whether their K reductase activity was maintained. The ratio of carboxylated FIXgla-PC produced in the KO-supplemented medium and the vitamin K-supplemented medium was used as a criterion to identify candidate cell colonies for VKOR gene knockout. To ensure that the knockout cell was clonal, functional screening was repeated as described earlier. To verify that the loss of KO reductase activity was due to TALENsmediated genome editing of the VKOR gene, genomic DNA was extracted from the selected VKOR knockout cells using QuickExtract DNA extraction solution (Epicentre, Madison, WI). The TALENs-targeted region in the genomic DNA was amplified by PCR using 50 GCTTCACTAGTCCCGGCATTCTTCGCTG-30 as the forward primer and 50 -CCAGCACTGTCTGGTCCCTTGCCTCGCAC-30 as the reverse primer. PCR products were cloned into the TOPO TA cloning vector (Invitrogen, Carlsbad, CA) for sequence analysis. To verify TALENs-mediated editing of the VKOR gene at the mRNA level, total RNA from VKOR knockout cells was extracted using Trizol. RT-PCR was performed using poly T18 and M-MLV reverse transcriptase (Invitrogen, Carlsbad, CA). The TALENs-targeted region was amplified

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by PCR, and the PCR product was cloned into the TOPO TA cloning vector for sequence analysis. It is worth noting that VKOR knockout cells regained KO reductase activity after culturing for several generations. It appears that this regained activity is from VKORL (Tie et al., 2013), a paralog enzyme of VKOR that shares
50% sequence identity with VKOR (Rost et al., 2004). Therefore, we designed TALENs pairs targeting the VKORL gene in the VKOR knockout cells. After loss-of-function screening, genomic DNA was prepared from the candidate knockout cell colonies and the targeted region was amplified by PCR using 50 -CCTTGCAATGTGCTAGTTAGACCGCT CC-30 as the forward primer and 50 -CACCAGGAAGACAGTGGGAAA GGCAACC-30 as the reverse primer. VKORL gene knockout has been confirmed by sequencing analysis of the target region. Cells with both VKOR and VKORL knocked out (DGKO) no longer support reporterprotein carboxylation using KO as the substrate (Fig. 10). However, exogenous expression of VKOR or VKORL significantly increases the production of carboxylated reporter protein, suggesting that the DGKO reporter cell line is a useful tool for VKOR function study with a clean background.

Fig. 10 Reporter protein carboxylation in the DGKO reporter cells. DGKO reporter cells were transiently transfected with expression vector (control), VKOR, or VKORL. Transfected cells were cultured with 5 μM KO for 48 h. The concentration of carboxylated FIXgla-PC in the cell culture medium was measured by ELISA.

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3.3 Functional Study of VKOR Using the DGKO Reporter Cells Functional study of VKOR is based on the ability of VKOR or its mutants to reduce KO to vitamin K to support the carboxylation of the reporter protein in the DGKO reporter cells. VKOR or VKOR mutant is transiently expressed in the DGKO cells, and the KO reductase activity is evaluated by measuring the level of carboxylated reporter protein in the cell culture medium when cells fed with KO. Mammalian expression vector pBudCE4.1 (Invitrogen, Carlsbad, CA), designed for simultaneous expression of two genes in mammalian cells, was used as the basic cloning vector. The cDNA of Metridia luciferase was cloned into one of the multicloning sites of pBudCE4.1 (the resulting plasmid is named as pBudCE4.1-Met.Luc) and the cDNA of VKOR or its mutant was cloned into the other multicloning site. Metridia luciferase, coexpressed with VKOR or its variants by the same vector controlled by different promoters, was used as the internal control for normalizing the transient transfection efficiency. For transient transfection, the DGKO cells were seeded, 1 day before transfection, into a 24-well plate in complete growth medium, so that the cells were approximately 60% confluent at the time of transfection. The plasmid DNA of pBudCE4.1-Met.Luc containing the cDNA of VKOR or its mutants was transiently transfected into the DGKO cells using Xfect transfection reagent (Clontech Laboratories, Inc., Mountain View, CA) according to the manufacturer’s instructions, with minor modifications. Triplet transfections were performed for each sample. For each well, 0.5 μg plasmid DNA was diluted with 25 μL of Xfect reaction buffer. After adding 0.15 μL of Xfect polymer, samples were mixed by vortexing for 10 s at high speed. Right before use, the Xfect polymer was diluted 20-fold with Xfect reaction buffer and mixed by vortexing. The DNA–Xfect polymer complex solution was incubated at room temperature for 10 min. Then the entire solution was added dropwise to one well containing 250 μL complete growth medium. The plate was gently rocked back and forth to mix and incubated at 37°C with 5% CO2 atmosphere for 4 h. The transfection medium was replaced with complete growth medium containing different concentrations of test compounds, such as KO alone or KO with different concentrations of warfarin. The plate was returned back to the 37°C CO2 incubator. Forty-eight hours posttransfection, the medium was collected to determine the concentration of carboxylated reporter protein and to measure luciferase activity for normalizing the transfection efficiency.

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The concentration of carboxylated reporter protein in the cell culture medium was measured by ELISA, as described in Section 2.3. Luciferase activity was determined by injecting 90 μL of 2 μM coelenterazine (NanoLight Technology, Pinetop, AZ) solution (in PBS with 300 mM NaCl) to 10 μL of cell culture medium in a 96-well Lumitrac white plate (Greiner Bio-One GmbH, Austria, Germany). Luminescence emission from the mixture was recorded using a SpectraMax L Luminescence Microplate Reader (Molecular Devices, Sunnyvale, CA) at 480 nm with an integration time of 1 s and a delay of 6 s. VKOR activity was evaluated by reporter-protein carboxylation normalized by luciferase activity for each individual sample. If a large number of samples need to be tested, the cell-based assays introduced in this chapter can be scaled up from the 24-well plate format to a 48-well plate format. In this case, 1 day prior to transfection, reporter cells would be seeded into 48-well plates in complete growth medium. An electronic adjustable tip spacing multichannel pipette is helpful for seeding the cells evenly in 48-well plates. For transfection, 0.25 μg plasmid DNA should be diluted with 12.5 μL Xfect reaction buffer. After mixing with Xfect polymer and incubating at room temperature for 10 min, the entire solution should be added dropwise to one well containing 125 μL complete growth medium. The plate should be rocked gently back and forth to mix, then incubated at 37°C with 5% CO2 atmosphere. After 4 h, instead of replacing the transfection medium with test medium, 125 μL of 2  test medium should be added directly to the transfection medium. The plate should be rocked gently back and forth to mix, and it should then be returned back to the 37°C CO2 incubator for 48 h.

3.4 Applications The established DGKO reporter cell line, which has all the essential endogenous components for VKD carboxylation except for KO reductase, provides an invaluable tool for studying the function of VKOR in its native milieu with a clean background. It is especially useful for studying the warfarin resistance of naturally occurring VKOR mutants (Fig. 11). Our knowledge on warfarin resistance of VKOR mutants is mainly based on in vitro studies that were carried out under artificial conditions using DTT to provide the reducing equivalents normally provided by unknown endogenous reductants of VKOR. Therefore, the warfarin resistance of most of the VKOR mutants as determined by the in vitro assay is not

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Fig. 11 Warfarin resistance study of VKOR and its Y139F mutant using the DGKO reporter cells. VKOR or its Y139F mutant was transiently expressed in the DGKO reporter cells and the transfected cells were cultured with 5 μM KO and increasing concentrations of warfarin for 48 h. The concentration of carboxylated FIXgla-PC in the cell culture medium was measured by ELISA.

consistent with the clinical phenotypes of patients carrying these mutations (Hodroge et al., 2012). Although the cell-based assay described in Section 2 has been successfully used for in vivo functional studies of VKOR, endogenous VKOR activity in HEK293 cells prevents its application for warfarin resistance studies, especially for those VKOR mutations with marginal resistance. Using the DGKO reporter cell line, we examined 10 naturally occurring VKOR mutants whose warfarin resistance could not be tested in vitro because they had essentially no in vitro activity (Hodroge et al., 2012). Results from the DGKO cell-based assay show that all these VKOR mutants are fully active (Tie et al., 2013). This is consistent with a recent clinical study showing that patients with warfarin-resistant VKOR mutations have normal VKD clotting factors in the absence of warfarin (Harrington, Siddiq, Allford, Shearer, & Mumford, 2011). Compared with the wild-type VKOR, five VKOR mutants showed increased warfarin resistance. These mutants, in increasing order, are: VKOR < S52L  N77S < W59L  L128R < W59R. Except for the L128R mutant, this order of warfarin resistance is consistent with the corresponding clinical anticoagulant dosages. However, the warfarin sensitivity of the other five VKOR mutants is not significantly different from that of the wild-type enzyme, which suggests that not all of the naturally occurring VKOR mutants found in patients requiring

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higher doses of warfarin are resistant to warfarin inhibition. Other factors must also play important roles in warfarin resistance in patients. Warfarin resistance in VKOR mutants has been studied using a nonknockout cell-based assay (Fregin et al., 2013), similar to the one described in Section 2. In general, the nonknockout cell-based assay reached a conclusion similar to that reached by the DGKO cell-based study. However, in the nonknockout cell-based assay, warfarin’s 50% inhibition (IC50) of both wild-type VKOR and the L128R mutant occurred at 24.7 and 1226.4 nM, respectively. These values are
10-fold higher than those obtained from the DGKO cell-based study (3.48 and 107 nM, respectively). The higher IC50 in the nonknockout cell-based assay is probably caused by endogenous VKOR in the reporter cells. An additional complication is that, instead of using KO as the substrate, this nonknockout cell-based assay used vitamin K as the substrate for the VKOR function study. Therefore, the VKOR activity measured by this approach depends on the conversion of vitamin K to KO by the HEK293 cells, a process that produces carboxylated reporter proteins independent of VKOR activity. The DGKO reporter cell line provides an ideal tool for studying VKOR function in a cellular environment. It is especially useful for studying the warfarin resistance of the naturally occurring VKOR mutations, and for clarifying VKOR’s genotype and the clinical phenotypes VKOR produces. Because of the depletion of VKORL in this cell line, it is also a potential tool for studying the structure–function relationship of VKORL, whose physiological role is still unknown.

4. FUNCTIONAL STUDY OF GGCX USING CRISPR-CAS9MEDIATED GENE KNOCKOUT REPORTER CELLS 4.1 Rationale The enzyme that accomplishes carboxylation is GGCX. It is responsible for carboxylation of VKD proteins to their functional forms. GGCX recognizes its protein substrate through a relatively tight binding to the propeptide of the VKD proteins, which tethers the substrate to the enzyme (Furie, Bouchard, & Furie, 1999). The binding of the propeptide to GGCX initiates a structural reorientation of GGCX by which the Gla domain of the protein substrate is positioned at the catalytic site of GGCX (HigginsGruber et al., 2010), and all 10–12 glutamic acids in the Gla domain are carboxylated during one binding event (Morris, Stevens, Wright, & Stafford, 1995). Although the apparent affinities of the VKD proteins’ propeptides

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for GGCX vary over 100-fold from in vitro studies (Higgins-Gruber et al., 2010), these VKD proteins appear to be fully carboxylated in physiological conditions. Genetic screening for GGCX variations in patients with vitamin K-related disorders has identified more than 30 naturally occurring mutations (Watzka et al., 2014). However, the correlation between these GGCX mutants and their clinical phenotypes is not clear. For example, it has been well established that mutations in the GGCX gene can result in defects of carboxylation of VKD coagulation factors causing bleeding disorders, referred to as VKCFD. The mainstay of therapy for VKCFD is the administration of vitamin K, which most often completely or partially corrects the coagulation factors defect; in some cases, however, vitamin K is ineffective (Brenner, Kuperman, Watzka, & Oldenburg, 2009). The patient’s vitamin K response does not seem to be correlated with either the clinical severity of the disorder or the severity of the molecular defect. Additionally, it is not clear why some mutations cause bleeding disorders while others cause nonbleeding syndromes such as Pseudoxanthoma elasticum (PXE)-like syndrome. This lack of knowledge is caused in part by the fact that our current understanding of GGCX’s function was obtained from in vitro experimentation under artificial conditions using the pentapeptide FLEEL as the substrate. Therefore, understanding GGCX, and being able to characterize this multiple-substrate enzyme in its native milieu, will help to clarify the clinical phenotypes and treatment of vitamin K-related disorders. In an attempt to access GGCX function in an animal model, the endogenous GGCX gene was knocked out in mice by gene targeting (Zhu et al., 2007). However, all homozygous GGCX-deficient mice succumbed to massive intraabdominal hemorrhage shortly after birth. Recently, in another attempt to study GGCX function in vivo, mice with liver-specific GGCX deficiency have been created (Azuma et al., 2014); however, manipulating GGCX variants and its substrates in the mouse model appears impracticable. Because manipulation of the vitamin K metabolism pathway is not lethal for mammalian cells, and because cell-based functional study of vitamin K cycle enzymes is feasible, we decided to develop a GGCX-deficient reporter cell line using CRISPR-Cas9-mediated genome editing to address questions related to GGCX catalysis in a cellular milieu. First, we stably expressed two structurally distinct VKD proteins (FIXgla-PT and MGP) as reporter proteins in HEK293 cells. FIXgla-PT is a chimeric protein of prothrombin with its Gla domain exchanged with that of factor IX. Like other VKD coagulation factors, prothrombin is

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produced and carboxylated in the liver. All the Gla residues of VKD coagulation factors, located in the N-terminal Gla domain, are preceded by the propeptide, which will be cleaved in the mature protein (Fig. 12). However, MGP, which is produced and carboxylated in extrahepatic tissues, retains its propeptide within the mature protein, and MGP’s Gla residues are scattered on both sides of the propeptide. While defects of carboxylation of coagulation factors result in bleeding disorders, defects of carboxylation of MGP are mainly associated with nonbleeding syndromes; we expect that these two structurally distinct reporter proteins may be carboxylated differently by GGCX, and that certain GGCX mutations could have different effects on these reporter proteins’ carboxylation, resulting in different clinical phenotypes. We used the genome-editing tool CRISPR-Cas9 to knock out the endogenous GGCX gene in HEK293 cells that stably express the two reporter proteins, FIXgla-PT and MGP. CRISPR-Cas9 mediates gene knockout by introducing targeted DSBs in the genomic DNA of the gene of interest (Cong et al., 2013; Mali et al., 2013). The introduction of the DSB is achieved by CRISPR-Cas9 with a specific single-guide RNA (sgRNA; Fig. 13A). The sgRNA binds to Cas9, directs it to a specific genomic locus complementary to the 20-base gRNA sequence, and induces a DSB using Cas9. Subsequent repair of the DSB via the NHEJ pathway in the cell produces indel mutations at the site of DSB. Frameshift mutations at the DSB site disrupt the gene of interest (Fig. 8B). Since GGCX is the only enzyme responsible for VKD carboxylation in the cell, knocking out of the endogenous GGCX gene will abolish reporter-protein carboxylation. Therefore, we can study GGCX function by observing how

Fig. 12 Domain structure of the reporter proteins MGP and FIXgla-PT. Top: Domain structure of MGP. MGP has a propeptide sequence in the mature protein. There is one Gla residue at the N-terminus and four Gla residues at the C-terminus of the propeptide sequence. Bottom: Domain structure of FIXgla-PT. Catalytic domain, region containing the serine protease catalytic triad of prothrombin; FIXgla, Gla domain of human factor IX; Kringle, regions of internal sequence homology of prothrombin; Propeptide, propeptide region of prothrombin.

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Fig. 13 Schematic diagram of CRISPR-Cas9-mediated gene knockout and optimized gRNA sequences for GGCX targeting. (A) Cas9-sgRNA in complex with a target site in the genomic DNA. sgRNA binds and directs Cas9 to the genomic DNA target site with the appropriate PAM sequence at the 30 end to create DSBs (indicated by the arrows). (B) Six optimized gRNA sequences for GGCX targeting and a nontargeting gRNA sequence used as the control. The gRNA sequence used for creating GGCX-knockout cell line is highlighted.

efficiently the reporter proteins are carboxylated in the GGCX-knockout cells by either the exogenously expressed GGCX or its mutants.

4.2 Establishment of the CRISPR-Cas9-Mediated GGCX Gene Knockout Reporter Cell Line To establish a reporter cell line with two reporter proteins (a dual-reporter cell line), FIXgla-PT and MGP were stably expressed in HEK293 cells. The DNA sequence encoding chimeric reporter-protein FIXgla-PT was created by overlap PCR, as previously described (Tie et al., 2011). The replacement in prothrombin of the Gla domain with the factor IX’s Gla domain allowed us to use the α-FIXgla mAb for quantitative detection of the carboxylated

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reporter protein. To increase reporter-protein stability, the protease cleavage sites at arginine residues 155, 271, and 320 of prothrombin were mutated to glutamine (Krishnaswamy, 2013). The cDNA of prothrombin with arginine 271 and 320 mutated to glutamine was a gift from Dr. Krishnaswamy from Children’s Hospital of Philadelphia, PA. MGP, the second reporter protein, has a C-terminal HPC4 tag (EDQVDPRLIDGK) for detection purposes. The DNA sequences encoding FIXgla-PT and MGP were cloned into the mammalian dual-expression vector pVITRO1-hygro-MCS (InvivoGen, San Diego, CA). Plasmid DNA of pVITRO1-hygro-MCS containing both sequences of FIXgla-PT and MGP was transfected into HEK293 cells. Forty-eight hours posttransfection, transfected cells were treated with trypsin, diluted with complete growth medium containing 0.3 mg/mL hygromycin, and seeded into 100-mm dishes. Single colonies were picked and screened for high expression of both reporter proteins. To knock out the endogenous GGCX gene of the above reporter cells, we used CRISPR-Cas9-mediated genome editing. First, we examined the genome-editing efficacy of six gRNA sequences optimized for GGCX targeting (Shalem et al., 2014) (Fig. 13B). Each of the six gRNA and a nontargeting gRNA (control) were cloned into the lentiCRISPR V2 vector (a gift from Feng Zhang, Addgene plasmid #52961; Shalem et al., 2014). This lentiCRISPR V2 vector allows the simultaneous expression of the sgRNA, endonuclease Cas9, and a puromycin selection marker. Lentivirus, produced by cotransfection of the lentiCRISPR V2 sgRNA constructs and the lentivirus packaging vectors (pVSVg and psPAX2) into HEK293T cells, was used to infect our reporter cells (FIXgla-PC/HEK293, Section 2.2) for the CRISPR-Cas9-mediated GGCX gene knockout. Twenty-four hours posttransduction, cells were cultured with complete medium containing 1.5 μg/mL puromycin; only cells transduced with a lentiCRISPR V2 sgRNA construct survived. Puromycin-resistant cells were expanded and cultured for 48 h with 11 μM vitamin K in the complete growth medium. For each gRNA, the efficiency of the genome editing in the GGCX knockout was evaluated by measuring the loss of reporter-protein carboxylation. The most efficient gRNA for GGCX targeting was used to knock out the endogenous GGCX gene in the dual-reporter cells. Next, the selected gRNA sequence (Fig. 13B) for GGCX targeting was cloned into plasmid pX330-U6-Chimeric-BB-CBh-hSpCas9 (a gift from Feng Zhang, Addgene plasmid #42230). This plasmid allows the simultaneous expression of the selected sgRNA, for GGCX targeting, and endonuclease Cas9, for making DSBs. It is transiently transfected into the

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dual-reporter cells. Forty-eight hours posttransfection, cells were treated with trypsin, and a single-cell suspension was obtained as described in Section 3.2. Diluted cell suspensions were plated into 96-well plates or seeded into 150-mm dishes for loss-of-function screening of GGCX knockout. To verify GGCX gene knockout in the candidate cell colonies, genomic DNA was prepared using QuickExtract DNA extraction solution. The gRNA-targeting region in the genomic DNA was amplified by PCR, and the PCR products were cloned into TOPO cloning vector for sequence analysis as described in Section 3.2. To further confirm the loss of endogenous GGCX expression in the GGCX-knockout cells, cell lysates were prepared and loaded to sodium dodecyl sulfate-polyacrylamide gel electrophoresis for western blot analysis. Protein bands were probed using rabbit anti-GGCX polyclonal antibody (Proteintech Group Inc., Chicago, IL). The selected positive GGCX-deficient cell colonies expressed no detectable endogenous GGCX and were unable to carboxylate the reporter proteins (Fig. 14). Expression of the exogenous GGCX in these GGCX-deficient cells restores carboxylation activity, suggesting that other components of the vitamin K cycle in the cells, as well as the reporter system, are intact.

4.3 Functional Study of GGCX Using the GGCX-Deficient Reporter Cells Functional study of GGCX is based on the ability of GGCX or its mutants to carboxylate the two reporter proteins in the GGCX-deficient dual-reporter cells. The cDNA of GGCX or its mutants was cloned into mammalian expression vector pBudCE4.1-Met.Luc and transiently expressed in the GGCX-deficient dual-reporter cells as described in Section 3.3. Four hours posttransfection, the transfection medium was replaced by complete growth medium containing 11 μM vitamin K and incubated for 48 h. Carboxylation activity was determined by measuring the level of carboxylated reporter proteins in the cell culture medium using sandwich-based ELISA (Tie et al., 2016). To evaluate carboxylation of FIXgla-PT, α-FIXgla mAb is used as the capture antibody, and horseradish peroxidase-conjugated sheep antihuman prothrombin (Affinity Biologicals Inc., Ancaster, Canada) is used as the detecting antibody. As described in Section 2.3, it is essential to include 3 mM Ca2+ in samples and all the buffers for ELISA. To measure the carboxylation of the second reporter protein MGP, a monoclonal antibody that specifically recognizes carboxylated MGP sequence 35 through 49 (VitaK

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Fig. 14 Functional characterization of the GGCX-knockout reporter cells. (A) Immunoblotting of wild-type (lane 1) and GGCX-knockout (lane 2) HEK293 cells. GGCX was probed by rabbit anti-GGCX polyclonal antibody (Proteintech Group Inc., Chicago, IL) as the primary antibody and horseradish peroxidase-conjugated goat antirabbit antibody (Jackson ImmunoResearch Laboratories, Inc, West Grove, PA) as the secondary antibody. The endogenous GGCX band is indicated by the asterisk. (B) Carboxylation activity of GGCX-knockout reporter cells and the cells transfected with wild-type GGCX. GGCX-knockout reporter cells were transiently transfected with expression vector (blank) or GGCX and cultured with 11 μM vitamin K for 48 h. The concentration of carboxylated FIXgla-PT in the cell culture medium was measured by ELISA.

BV, Maastricht, the Netherlands) is used as the capture antibody, and biotinylated anti-HPC4 monoclonal antibody is used as the secondary antibody. Streptavidin poly-HRP (Pierce, Rockford, IL) is used to amplify the signal of MGP detection. For MGP detection, it is important to not include Ca2+ in sample and all the buffers for ELISA.

4.4 Applications The established GGCX-deficient dual-reporter cell line has been successfully used to study GGCX function and to clarify the correlation between GGCX genotypes and their clinical phenotypes. Using this cell-based assay, we first studied GGCX mutations found in a child with severe cerebral bleeding and comorbid Keutel syndrome, a nonbleeding malady caused by functional defects of MGP (Tie et al., 2016). Clinical data showed that high doses of vitamin K administration improved the bleeding diathesis of the patient, but not the functional defect of MGP. Genetic analysis of the

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GGCX gene in the patient detected three missense mutations corresponding to amino acid changes of D153G, M174R, and R325Q. To study how these GGCX mutations impacted the clinical phenotypes of the patient, we measured the carboxylation activity of these mutants using the GGCX-deficient dual-reporter cell-based assay. Results showed that the R325Q mutant is almost as efficient as the wild-type enzyme for the carboxylation of both reporter proteins. In contrast, the M174R mutant carboxylates neither reporter protein in the presence of vitamin K. As the patient had a defect of VKD coagulation factors—a defect that can be ameliorated by vitamin K supplementation—these two mutations are unlikely to explain the patient’s clinical phenotypes. However, while the D153G mutant has
60% activity for FIXgla-PT carboxylation in the presence of high concentration of vitamin K, it displays significantly decreased MGP carboxylation. This result is consistent with the patient’s clinical result, suggesting that D153G is the causative mutation for the patient’s clinical phenotypes. To gain further insights on how the clinical phenotypes respond to vitamin K administration, GGCX-deficient dual-reporter cells expressing the causative GGCX mutant D153G were treated with increasing concentrations of vitamin K. Results show that higher vitamin K concentrations recovered the FIXgla-PT carboxylation up to
60% (Fig. 15A). This is

Fig. 15 Vitamin K titration of GGCX and its D153G mutant using GGCX-knockout reporter cells. Carboxylation of reporter proteins FIXgla-PT (A) and MGP (B) by GGCX (•) and D153G mutant (■) with increasing concentrations of vitamin K. GGCX or its D153G mutant was transiently expressed in the GGCX-knockout reporter cells and the transfected cells were cultured with increasing concentrations of vitamin K for 48 h. The concentrations of carboxylated reporter proteins in the cell culture medium were measured by ELISA.

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consistent with the patient’s clinical results, in which the VKD coagulation factor’s activity reached up to 53% after supplementation with high-dose vitamin K. Additionally, the cell-based assay shows that higher concentrations of vitamin K have only a moderate effect on MGP carboxylation (Fig. 15B), a finding that again agrees with the clinical results of the patient; administration of high doses of K vitamins failed to increase the carboxylation of MGP. These results, taken together, suggest that the GGCXdeficient dual-reporter cell-based assay is a good system for the functional study of GGCX in vivo. One of the caveats of using results obtained from the GGCX-deficient cell-based assay to explain clinical phenotypes is that two of the reporter proteins in this assay are carboxylated in the same type of cell, and factors that affect VKD proteins’ carboxylation in physiological conditions may behave differently in different types of cells/tissues. For example, vitamin K (phylloquinone) is the main supplement used to treat VKCFD patients to improve the carboxylation of coagulation factors. It is the most abundant form of K vitamin in the liver (Shearer & Newman, 2014), where coagulation factors are produced and carboxylated. On the other hand, administration of menaquinones, but not phylloquinone, improves MGP carboxylation and reduces both cardiovascular mortality and coronary calcification. MGP is produced by chondrocytes, vascular smooth muscle cells, endothelial cells, and fibroblasts, and it undergoes local carboxylation in the extrahepatic tissues, where menaquinones are believed to play a role in VKD carboxylation (Danziger, 2008). Menaquinone supplementation has been shown to improve extrahepatic vitamin K status, but to have no effect on thrombin generation in healthy subjects (Theuwissen et al., 2012). However, results from the GGCX-deficient cell-based assay show that both vitamin K and menaquinones can efficiently support the carboxylation of both coagulation factors and MGP by wild-type GGCX (Tie et al., 2016). The different contributions of phylloquinone and menaquinones to hepatic and extrahepatic carboxylation may therefore be caused by the way different tissue absorbs and utilizes these K vitamins.

5. CONCLUDING REMARKS The cell-based assays introduced in this chapter provide versatile tools for the functional study of enzymes of the vitamin K cycle in a cellular milieu. With the aid of recently developed genome-editing techniques, one can knock out a specific gene of interest in the reporter cell line and

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study the function of the protein in the native cellular environment. We expect that these novel cell-based assays will improve our understanding of how the various vitamin K cycle components contribute to these proteins’ complex mechanisms and functions. The detailed understanding of VKOR and GGCX and their naturally occurring mutants that can be achieved using these assays will lead to new ways of controlling thrombosis, will help researchers understand warfarin therapy, and will aid in the development of new anticoagulant drugs. In addition, it is becoming clear that VKD proteins control physiologic processes that contribute to other areas of health interest. Therefore, it seems likely that studies using these cell-based assays will lead to a better understanding of the mechanisms of a variety of diseases.

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Tokunaga, F., Takeuchi, S., Omura, S., Arvan, P., & Koide, T. (2000). Secretion, gammacarboxylation, and endoplasmic reticulum-associated degradation of chimeras with mutually exchanged Gla domain between human protein C and prothrombin. Thrombosis Research, 99(5), 511–521. Tokunaga, F., Wakabayashi, S., & Koide, T. (1995). Warfarin causes the degradation of protein C precursor in the endoplasmic reticulum. Biochemistry, 34(4), 1163–1170. Ulrich, M. M., Furie, B., Jacobs, M. R., Vermeer, C., & Furie, B. C. (1988). Vitamin K-dependent carboxylation. A synthetic peptide based upon the gamma-carboxylation recognition site sequence of the prothrombin propeptide is an active substrate for the carboxylase in vitro. The Journal of Biological Chemistry, 263(20), 9697–9702. Vermeer, C., Soute, B. A., De Metz, M., & Hemker, H. C. (1982). A comparison between vitamin K-dependent carboxylase from normal and warfarin-treated cows. Biochimica et Biophysica Acta, 714(2), 361–365. Vermeer, C., Soute, B. A., Hendrix, H., & de Boer-van den Berg, M. A. (1984). Decarboxylated bone Gla-protein as a substrate for hepatic vitamin K-dependent carboxylase. FEBS Letters, 165(1), 16–20. Wajih, N., Hutson, S. M., Owen, J., & Wallin, R. (2005). Increased production of functional recombinant human clotting factor IX by baby hamster kidney cells engineered to overexpress VKORC1, the vitamin K 2,3-epoxide-reducing enzyme of the vitamin K cycle. The Journal of Biological Chemistry, 280(36), 31603–31607. http://dx.doi.org/10.1074/ jbc.M505373200. Wajih, N., Hutson, S. M., & Wallin, R. (2007). Disulfide-dependent protein folding is linked to operation of the vitamin K cycle in the endoplasmic reticulum. A protein disulfide isomerase-VKORC1 redox enzyme complex appears to be responsible for vitamin K1 2,3-epoxide reduction. The Journal of Biological Chemistry, 282(4), 2626–2635. http:// dx.doi.org/10.1074/jbc.M608954200. Wallin, R., Gebhardt, O., & Prydz, H. (1978). NAD(P)H dehydrogenase and its role in the vitamin K (2-methyl-3-phytyl-1,4-naphthaquinone)-dependent carboxylation reaction. The Biochemical Journal, 169(1), 95–101. Wallin, R., & Martin, L. F. (1987). Warfarin poisoning and vitamin K antagonism in rat and human liver. Design of a system in vitro that mimics the situation in vivo. The Biochemical Journal, 241(2), 389–396. Wallin, R., Sane, D. C., & Hutson, S. M. (2002). Vitamin K 2,3-epoxide reductase and the vitamin K-dependent gamma-carboxylation system. Thrombosis Research, 108(4), 221–226. Wang, X., Dutton, R. J., Beckwith, J., & Boyd, D. (2011). Membrane topology and mutational analysis of Mycobacterium tuberculosis VKOR, a protein involved in disulfide bond formation and a homologue of human vitamin K epoxide reductase. Antioxidants & Redox Signaling, 14(8), 1413–1420. http://dx.doi.org/10.1089/ars.2010.3558. Watzka, M., Geisen, C., Scheer, M., Wieland, R., Wiegering, V., Dorner, T., … Oldenburg, J. (2014). Bleeding and non-bleeding phenotypes in patients with GGCX gene mutations. Thrombosis Research, 134(4), 856–865. http://dx.doi.org/10.1016/j. thromres.2014.07.004. Westhofen, P., Watzka, M., Marinova, M., Hass, M., Kirfel, G., Muller, J., … Oldenburg, J. (2011). Human vitamin K 2,3-epoxide reductase complex subunit 1-like 1 (VKORC1L1) mediates vitamin K-dependent intracellular antioxidant function. The Journal of Biological Chemistry, 286(17), 15085–15094. http://dx.doi.org/10.1074/jbc.M110.210971. Whitlon, D. S., Sadowski, J. A., & Suttie, J. W. (1978). Mechanism of coumarin action: Significance of vitamin K epoxide reductase inhibition. Biochemistry, 17(8), 1371–1377. Willems, B. A., Vermeer, C., Reutelingsperger, C. P., & Schurgers, L. J. (2014). The realm of vitamin K dependent proteins: Shifting from coagulation toward calcification.

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CHAPTER FIFTEEN

Activity Assays for Rhomboid Proteases E. Arutyunova*, K. Strisovsky†, M.J. Lemieux*,1 *Faculty of Medicine and Dentistry, Membrane Protein Disease Research Group, University of Alberta, Edmonton, AB, Canada † Institute of Organic Chemistry and Biochemistry, Academy of Sciences of the Czech Republic, Prague, Czech Republic 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. In Vivo Assays 2.1 In Bacteria 2.2 In Mammalian Cells 3. In Vitro Assays 3.1 Recombinant Transmembrane Protein Substrates and Gel-Based Readout 3.2 Fluorescent Substrates Acknowledgments References

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Abstract Rhomboids are ubiquitous intramembrane serine proteases that are involved in various signaling pathways. This fascinating class of proteases harbors an active site buried within the lipid milieu. High-resolution structures of the Escherichia coli rhomboid GlpG with various inhibitors revealed the catalytic mechanism for rhomboid-mediated proteolysis; however, a quantitative characterization was lacking. Assessing an enzyme’s catalytic parameters is important for understanding the details of its proteolytic reaction and regulatory mechanisms. To assay rhomboid protease activity, many challenges exist such as the lipid environment and lack of known substrates. Here, we summarize various enzymatic assays developed over the last decade to study rhomboid protease activity. We present detailed protocols for gel-shift and FRET-based assays, and calculation of KM and Vmax to measure catalytic parameters, using detergent solubilized rhomboids with TatA, the only known substrate for bacterial rhomboids, and the model substrate fluorescently labeled casein.

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1. INTRODUCTION Rhomboids are ubiquitous intramembrane serine proteases that are involved in various signaling pathways (Strisovsky, 2016; Urban & Dickey, 2011). This fascinating class of proteases harbors an active site buried within the lipid milieu. High-resolution structures of the Escherichia coli rhomboid GlpG alone and with various inhibitors and peptides revealed the catalytic mechanism for rhomboid-mediated proteolysis (Cho, Dickey, & Urban, 2016; Lemieux, Fischer, Cherney, Bateman, & James, 2007; Vinothkumar, 2011; Vinothkumar et al., 2011; Vosyka et al., 2013; Wang, Zhang, & Ha, 2006; Zoll et al., 2014); however, quantitative characterization of activity was needed to provide a thorough understanding. Assessing an enzyme’s catalytic parameters is important for understanding the details of its proteolytic reaction and regulatory mechanisms. To assay rhomboid protease activity, many challenges exist. Rhomboid proteases are embedded within lipid bilayers and cleave transmembrane substrates, making analysis complicated since detergents are needed to extract the protein from the membrane. Also, isolation of both enzymes and substrates is complex due to their high hydrophobicity and sensitivity to denaturation. Importantly, a lack of known substrates has impeded analysis of rhomboid protease and in particular the E. coli rhomboid protease GlpG, which has served as a model for the rhomboid protease family. The first activity assays for rhomboid proteases employed model transmembrane protein substrates and SDS-PAGE-based readout (Lemberg et al., 2005; Maegawa, Ito, & Akiyama, 2005; Strisovsky, Sharpe, & Freeman, 2009; Urban, Schlieper, & Freeman, 2002; Urban & Wolfe, 2005). While this is an accurate and reproducible assay format, it is also low-throughput, relatively time-consuming, and allows only an endpoint analysis of the reaction. An interesting variation of this assay employs detection and quantitation of reaction products by mass spectrometry (Vosyka et al., 2013), which, however, requires an expensive instrument for detection and is not continuous, limiting its practical use. Substrates that allow continuous assays and measurement of initial reaction rates using optical readout are much more practical, and will be the main focus of this article. A generic fluorogenic protease substrate, internally quenched BODIPYcasein, can be cleaved by GlpG (Maegawa et al., 2005; Wang et al., 2006), although it is a completely heterologous substrate lacking a transmembrane domain, and thus does not mimic the interactions of a true rhomboid

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substrate. More native and advanced assays that enable measurement of enzyme kinetics use fluorogenic peptide or protein substrates: a Forster resonance energy transfer (FRET) derivative of the truncated Gurken substrate (Pierrat et al., 2011), a genetically encoded transmembrane FRET substrate based on a CyPet-TatA-YPet recombinant fusion protein (Arutyunova et al., 2014), or fluorescein isothiocyanate (FITC)-labeled TatA transmembrane peptide (Dickey, Baker, Cho, & Urban, 2013). Although these assays are currently the most advanced ways to measure rhomboid protease kinetics, each of these assays has its limitations that would manifest during largescale screening for inhibitors. Some of these assays work only for a specific rhomboid protease (Pierrat et al., 2011), while others are usable only in liposomes (Dickey et al., 2013). An alternative tool for detection of proteases and their inhibitors is activity-based probes (Bachovchin, Brown, Rosen, & Cravatt, 2009). A fluorescence polarization assay with activity-based probes was used to identify new β-lactone-type inhibitors (Wolf et al., 2013). The significant advantage of these phosphonofluoridate probes is that they are relatively unselective, and might be usable for virtually any rhomboid protease. On the other hand, fluorescence polarization is prone to detergent artifacts, and an active-site-directed probe would most likely not detect an allosteric inhibitor or an activator. In this chapter, we give the reader an overview of substrate cleavage-based activity assays for rhomboid proteases with particular emphasis on in vitro gel-based assays and continuous, fluorescence-based assays.

2. IN VIVO ASSAYS 2.1 In Bacteria The activity of bacterial rhomboid proteases has been measured in live cells using a number of substrate constructs. Akiyama et al. used a LacYTM2based substrate to analyze substrate specificity of the endogenous GlpG in E. coli (Akiyama & Maegawa, 2007; Maegawa et al., 2005), Spitz transmembrane domain (TMD)-based substrate was used by Urban et al. using coexpression with recombinant rhomboid (Baker, Young, Feng, Shi, & Urban, 2007; Moin & Urban, 2012; Urban & Baker, 2008; Urban & Wolfe, 2005), and Strisovsky et al. constructed a chimeric framework consisting of maltose-binding protein (MBP), substrate transmembrane domain, and thioredoxin, in which a number of wild-type and mutant transmembrane domains were used in specificity studies (LacYTM2, Spitz,

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Gurken, TatA, and EGF) (Adrain et al., 2011; Strisovsky et al., 2009). The TatA protein and its cognate rhomboid AarA from Providencia stuartii were used in a large mutagenesis study that defined rhomboid proteases recognize two elements in their substrates, one of which is a linear sequence motif spanning approximately the P4 to P2ʹ positions (Strisovsky et al., 2009). All of these in vivo assays are endpoint measurements that use SDS-PAGE as a readout. These assays have been adapted for the measurement of inhibition of GlpG by using an E. coli strain with a mutant outer membrane protein (Ruiz, Falcone, Kahne, & Silhavy, 2005), which increases the permeability of the outer membrane and improves access of small molecules into the periplasm of E. coli (Cho et al., 2016; Pierrat et al., 2011).

2.2 In Mammalian Cells The activity of eukaryotic rhomboid proteases has traditionally been measured in cultured animal cells, because they are generally much more difficult to prepare in recombinant purified form. Plasmid-encoded epitope-tagged rhomboids were coexpressed with their epitope-tagged substrates in mammalian cells (COS1, COS7, or HEK293T) by transient transfection, and the intracellular cleavage of substrates or secretion of their ectodomains into the media were analyzed by Western blotting (Adrain et al., 2011; Baker, Wijetilaka, & Urban, 2006; Lemberg et al., 2005; Lohi, Urban, & Freeman, 2004; Urban, Lee, & Freeman, 2001). At present this is the only practical way to measure the activity and inhibition of some eukaryotic rhomboid proteases, but the fact that it is based on SDS-PAGE and Western blotting makes it unsuitable for large-scale screening of inhibitors. An improved, higher throughput assay for rhomboid activity in mammalian cells would be very useful.

3. IN VITRO ASSAYS A continuous in vitro assay using purified components is the most direct reporting system for protease activity, but it requires that enzymes be purified in a state with retained activity, which is not always easy to achieve. Furthermore, substrates also need to be purified to homogeneity to allow for proper measurement of catalytic parameters. As mentioned earlier, two main methods that have been developed for in vitro kinetic analysis use either detergent solubilized or lipoprotein reconstituted rhomboid proteases. Both have limitations. For example, detergent solubilization, while stripping most lipids from membrane proteins may compromise structure

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and function. Rhomboid proteases reconstituted into proteoliposomes require reconstitution with substrate and coordination of cleavage site with active site in the membrane during reconstitution. The readout for these assays involves SDS-PAGE or fluorescence measurement depending on the substrates used.

3.1 Recombinant Transmembrane Protein Substrates and Gel-Based Readout One of the first attempts to characterize functional features of rhomboids before any of prokaryotic substrate was known was the development of a model substrate based on N-terminal and periplasmically localized β-lactamase (Bla) domain, a LacY-derived transmembrane region, and a cytosolic MBP mature domain (Maegawa et al., 2005). The distinctive feature of this substrate was its precise design to mimic the natural properties of known rhomboids substrates—the fusion protein was engineered to have the type I (Nout–Cin) topology and the cleavage site was located in the transmembrane domain. Using Bla-LacYTM2-MBP-His6 model membrane substrate, Akiyama group established gel-shift in vitro and in vivo assays for assessing proteolytic activity of E. coli rhomboid. Interestingly, it was found that the cleavage site for rhomboids was located in the region with high local hydrophilicity. Another substrate used in gel-based rhomboid assay was adapted from the assay for best understood intramembrane protease at that time—γ-secretase (Li et al., 2000). To convert a γ-secretase substrate into a rhomboid substrate, the first seven residues of a C-terminally Flag-tagged C100, a recombinant C-terminal fragment of Alzheimer’s amyloid precursor protein that expresses to very high levels, were replaced by the first seven residues from Spitz, a Drosophila melanogaster rhomboid substrate. This substitution was necessary and sufficient to convert C100-Flag protein into substrate for rhomboids (Urban & Freeman, 2003). The activity assay developed with this substrate demonstrated for the first time that rhomboid proteases do not require cofactors to catalyze the cleavage reaction; they directly recognize their substrate, and their activity is affected by the presence of lipids (Urban & Wolfe, 2005). A later version of the assay, using the same substrate, was used to investigate the gating mechanism of E. coli rhomboid (Baker et al., 2007). In order to test the cleavage of Spitz in bacteria with coexpressed of GST-rhomboids another substrate was designed—the chimeric protein was composed of bacterial pelB leader peptide to target the protein to the membrane, green fluorescent protein (GFP) as the

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extracellular ectodomain, the juxtamembrane–transmembrane–cytosolic residues (123–230) of Spitz, and a C-terminal flag epitope. The recombinant expression of this substrate with Bacillus subtilis YqgP and P. stuartii AarA rhomboids in E. coli cells demonstrated strong cleavage (Urban & Wolfe, 2005). The Freeman group developed chimeric substrates based on transmembrane domains of four known model substrates of bacterial rhomboid proteases: TMD of P. stuartii TatA (Stevenson et al., 2007), TMD2 of E. coli LacY (Maegawa et al., 2005), and D. melanogaster Gurken and Spitz (Lemberg et al., 2005; Urban, Lee, & Freeman, 2002). These substrates constructs included a signal peptide flanked by N-terminal MBP and C-terminal thioredoxin with His-tag (Strisovsky et al., 2009). Three rhomboids [AarA (P. stuartii), GlpG (E. coli), and YqgP (B. subtilis)] were used to analyze the cleavage of the recombinant substrates. The presence of fusion proteins did not affect the cleavage of TMD by rhomboids; the same phenomenon we also observed for psTatAFRET-substrate. The recombinant substrates were also generated by in vitro translation in the presence of L-[35S]-Met to radiolabel them, which facilitates cleavage detection. Using these chimeric substrates it was demonstrated that rhomboids recognize a specific sequence surrounding the cleavage site and their cleavage event is sequence-specific (Strisovsky et al., 2009). Natural substrates have also been used in in vitro assays but with limitations. Low expression yields limits analysis of eukaryotic rhomboids and their natural substrates. For bacterial rhomboid proteases, we are constrained in our knowledge by only one physiological substrate, namely the TatA translocon protein from P. stuartii cleaved by the rhomboid AarA. Our group developed an assay to assess the catalytic parameters of cleavage of AarA rhomboid protease with its natural substrate using a gel-based readout. The full-length P. stuartii TatA (psTatA) gene was expressed and purified from pBad-Myc-HisA plasmid construct (Arutyunova et al., 2014). A good separation on the gel between a substrate band and the product band is a key for precise velocity determination. However, the cleavage site for rhomboid is located too close to the N-terminus of the protein. Therefore, in order to increase the size difference between uncleaved and cleaved psTatA a small linker as well as Flag-tag was introduced at the N-terminus (for purification and detection purposes), which is a similar strategy used previously (Strisovsky et al., 2009). This assay system will now be described in more detail. In order to perform this type of assay, a selective method able to distinguish between the cleaved and uncleaved substrate is required. In most cases,

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electrophoresis is used to separate the product from undigested substrate. In order to stop the reaction, enzyme denaturation conditions such as SDSsample buffer can be used. Although this type of assay is amenable to higher throughput, there is a high probability of introducing timing and volume inaccuracies. In addition, this method does not reveal the dependence of product formation on time, which is essential for accurate and advanced kinetics measurements. Therefore, several parameters need to be checked to ensure the validity of discontinuous assay. The underlying assumption of any discontinuous assay is that the sample is taken within the initial velocity (linear) phase of the reaction, and therefore the product formation is a linear function of time. If this condition is met then from the time course of product formation plot, the initial velocity can be expressed as: vo ¼ ΔS=Δt, where ΔS is the change in substrate concentration that occurred during time Δt. If the signal at zero time point is negligible or equals zero then the velocity equals S/t. Thus, if the time interval is fixed then the product signal becomes proportional to velocity and can be used for conducting saturation plots. However, experimental verification is needed to prove that the chosen time point is within the linear part of the reaction curve. Therefore, prior the assay several parameters need to be optimized. The following sections will describe in detail the protocols for expression and purification of psTatA substrate, optimization of assay parameters, and the kinetics analysis of rhomboid catalysis. 3.1.1 Expression and Purification of psTatA Substrate 1. Transform pBad-Myc-His vector, bearing psTatA gene into K12 GlpG knockout cells competent cells (glpG::Kn) from Keio library and select transformants on LB plates containing 100 μg/mL of ampicillin. The E. coli Keio knockout collection contains a set of single-gene, knockout mutants for all nonessential genes in E. coli K-12 (Baba et al., 2006). We use these cells in order to eliminate the cleavage of expressed psTatA by endogenous GlpG. Even though psTatA is not a native substrate for E. coli rhomboid, it has been demonstrated that GlpG is able to cleave psTatA (Arutyunova et al., 2014; Lazareno-Saez, Arutyunova, Coquelle, & Lemieux, 2013; Strisovsky et al., 2009). 2. Pick a single transformant colony and inoculate into 150 mL of LB medium, supplemented with 100 μg/mL of Amp (LB-Amp). Shake

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overnight (200 rpm) at 37°C. The following day, subinoculate 20 mL of the overnight culture into three individual 4 L flasks containing 1 L of LB-Amp. Grow cells at 37°C until OD600 reaches 0.6, induce the protein with 0.02% arabinose, and express at 24°C for 6 h. After harvesting the cell culture, lyse the cells and isolate the membrane fraction, conducting two subsequent centrifugation runs: low speed at 20,000  g for 25 min to remove unbroken cells and cell debris and high speed at 100,000  g for 2 h to pellet the membranes. Homogenize the membrane fraction in 1:10 (weight:volume) ratio of 50 mM Tris–HCl, pH 8.0, 200 mM NaCl, 20% glycerol buffer, and solubilize the membrane proteins with 1% DDM for 30–60 min. Remove unsoluble particles by high-speed centrifugation at 100,000  g for 30 min. Apply the supernatant from previous step on Anti-Flag M2 Affinity Gel (Sigma-Aldrich, USA). Elute psTatA from the matrices with 0.1 M glycine, pH 3.5, 0.1% DDM, and adjust the pH immediately with 1 M Tris–HCl, pH 7.5, 0.1% DDM. Analyze the purity of protein with SDS-PAGE, assess the concentration of protein sample with the Pierce BCA protein assay kit (Thermo Fisher, USA), concentrate using Amicon 3 kDa MWCO filtration device (EMD Millipore, USA), and flash-freeze immediately. The bicinchoninic acid (BCA) protein assay method (Smith et al., 1985) is compatible with certain amounts of detergents. As it is shown in Fig. 1, Anti-Flag resin allows obtaining pure protein in only one purification step.

Fig. 1 SDS-PAGE of psTatA purification. Lane 1: protein markers, lanes 2–4: fractions from Anti-Flag M2 resin.

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3.1.2 Optimization of Rhomboid Protease Concentration and the Reaction Time In order to ensure the assay validity several parameters as enzyme concentration and time of reaction need to be optimized. For discontinuous assays it is important to stop the reaction at various times of incubation to determine the enzyme concentration, which gives the linear rate throughout a certain period of time and consequently to define the time of reaction. 1. Make a serial dilutions of rhomboid protease using activity buffer (50 mM MES, pH 6.0, 150 mM NaCl, 20% glycerol, 0.1% DDM), so the final enzyme concentrations in the reaction mixture is 0.05, 0.25, and 1.5 μM. 2. Prepare the activity reaction mixtures containing 10 μM of psTatA substrate and the activity buffer. The reaction mixture should be a sufficient volume, so that the convenient size samples can be removed, quenched, and analyzed at different time points throughout the reaction. The concentration of substrate used for this experiment should be high enough not to change during the reaction time, otherwise when the substrate concentration is too low ([S]  KM) reaction rate decreases, which impedes initial rate determination. Preincubate the substrate with the activity buffer at 37°C for 30 min. Preincubation time is necessary for the system to reach equilibrium so the system in steady-state condition and its behavior does not change in time. In many cases, a steady state is not achieved until sometime after the system is started. 3. Start the series of reactions with the equal volume of rhomboid’s dilutions. 4. Remove an aliquot of the reaction mixture after 30 min, 1, 2, 3, 5, and 6 h. Immediately after sample removal add the equal volume of 4SDSsample buffer. The increased concentration of SDS-sample buffer is required to ensure that the condition for quenching of reaction lead to instantaneous and permanent halt of signal production. This is very important condition of endpoint assay, because if the reaction is not stopped but just slowed down, serious problems with assay reproducibility may be encountered later. After addition of SDS-sample buffer the samples should be put on ice or frozen if the analysis is not performed immediately. 5. The reaction mixture with the same volume and amount of substrate should be used as negative control. The samples should be removed at the same time points as for reaction samples, proving the stability of the substrate during incubation time at 37°C.

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6. Load the equal amount of samples on Tricine-SDS gel. We utilized Tricine-SDS-PAGE instead of regular Laemmli-SDS-PAGE to improve the separation of the bands. Tricine-SDS-PAGE is the preferred electrophoretic system for the resolution of proteins smaller than 30 kDa. It is also used for the samples containing hydrophobic proteins of interest that should be transferred onto a membrane for Western blot detection (Schagger, 2006). Run the gel at 30 V for stacking gel and at 100 V for resolving gel to attain good separation. 7. Perform the Western blot: transfer the bands onto nitrocellulose membrane and reveal the protein with 6His epitope tag antibodies (Thermo Fisher Scientific, USA). 8. Quantify the bands by densitometry measurement, which in our case was done with ImageQuant LAS 4000 (GE Healthcare, USA). Densitometry measurement is a result of the darkness of a given area and then is expressed as a number of a dark spots and in a relative mean gray value (MGV). 9. Plot obtained MGVs vs time for each enzyme concentration. Fig. 2 illustrates the correlation between the concentration of enzyme and time of reaction. In case of 0.05 μM of rhomboid, the rate is too slow and even if it shows linear proportionality the initial velocity is understated. In the

Fig. 2 The time course of psTatA cleavage by different concentrations of AarA.

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curve with 1.5 μM of rhomboid protease the presence of burst indicates that the concentration of enzyme is too high. The reaction rate with 0.25 μM of rhomboid shows a long linear phase with the plateau phase after 5 h. Thus, the time of reaction up to 4 h and 0.25 μM of enzyme are the optimal conditions for the kinetic assay of AarA-mediated cleavage of psTatA.

3.1.3 Proportionality Between Chemiluminescent Signal and the Concentration of Substrate To quantify substrate and product, Western blot can be used with chemiluminescence detection methods. There are several parameters one should be aware of when performing kinetic assay that involves the chemiluminescence readings—sensitivity, linearity, and dynamic range. Sensitivity refers to the lowest concentration of substrate that will be reliably detected. This concentration will set the lowest substrate concentration limit for the assay. Linearity describes the relationship of chemiluminescence signal to the amount of substrate over a range of concentrations used in the assay. Ideally, the plot of signal vs substrate should be linear. Dynamic range is a span of substrate concentrations, which can be detected at the same time without losing proportionality. This sets the working range of the assay. 1. Make serial dilutions of psTatA protein in the activity buffer (50 mM MES, pH 6.0, 150 mM NaCl, 20% glycerol, 0.1% DDM) where the final concentrations match the final concentrations of psTatA used in the assay (0.5, 1, 2, 3, 4, 5, 7, 10, and 15 μM). 2. Add equal volume of 4SDS-sample buffer to the protein sample. 3. Load 20 μM of protein sample on 16% Tricine-SDS gel. 4. Perform the Western blot by transferring the bands onto nitrocellulose membrane and revealing the protein with 6His epitope tag antibodies (Thermo Fisher Scientific, USA) (Fig. 3). 5. Quantify the bands by densitometry measurement, which in our case was done with ImageQuant LAS 4000 (GE Healthcare, USA). 6. Plot MGVs vs concentration of psTatA. As demonstrated in Fig. 3, the dependence of chemiluminescent signal on psTatA concentration represents a straight line. Thus, the concentration range used for kinetic assay of AarA-mediated cleavage is valid. Proportionality check is also important because when broad concentration range is used, the substrate conversion at low substrate concentrations is high, which may lead to underestimation of initial reaction rates (Purich, 2010).

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Fig. 3 The proportionality between chemiluminescence and concentration of psTatA. (A) Western blot of increasing concentrations of psTatA, visualized with 6His epitope tag antibody. (B) The dependence of chemiluminescence signal (MGV) of different psTatA concentrations on concentration.

3.1.4 Kinetic Analysis of AarA-Mediated Cleavage of psTatA 1. Prepare reaction mixtures containing psTatA in the concentration range from 0.5 to 15 μM and activity buffer (50 mM MES, pH 6.0, 150 mM NaCl, 20% glycerol, 0.1% DDM). Incubate the reaction mixtures at 37°C for at least 30 min to allow the system to equilibrate. 2. Start the reaction with the addition of 0.25 μM of AarA rhomboid. 3. Incubate the samples for 2 h at 37°C. 4. The samples with the same concentrations of substrate but with no protease should be incubated at 37°C for 2 h to be used as negative controls in order to ensure that there is no degradation after the time of reaction. 5. Stop the reactions by adding the same volume of 4SDS-sample buffer. 6. Reveal protein samples with Western blot using 6His epitope tag antibodies (Thermo Fisher Scientific, USA).

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7. Quantify the bands by densitometry measurement. Estimate the percentage of cleaved substrate, obtained from MGVs of digested and undigested substrates. Product ð%Þ ¼ MGV ðdigestedÞ  100=ðMGV ðdigestedÞ + MGV ðundigestedÞÞ 8. Calculate the amount of generated product using the known initial concentration of substrate in μM. 9. Plot the generated product vs substrate concentration. For AarA, we fit the plot with both Michaelis–Menten and the Hill equations using Prism software (GraphPad, USA) (Fig. 4). Choose the preferred fit based on R2 values and analysis of the residuals; these parameters are

Fig. 4 (A) Western blot analysis of AarA-mediated cleavage of psTatA. psTatA at the concentration range from 0.5 to 15 μM was incubated with 0.25 μM of AarA at 37°C for 2 h; the cleavage product (C) was separated from the uncleaved substrate (U) with SDS-PAGE and developed with Western blot using 6His epitope tag antibodies. The star represents a contaminant band from the substrate sample. (B) Hill plot of formation of AarA-cleaved psTatA vs psTatA concentration (n ¼ 2, mean  SD). The catalytic parameters of cleavage were calculated from Hill equation: K0.5 ¼ 7.6  1.1 μM, kcat ¼ 1.06  0.05 min–1, Hill coefficient ¼ 1.7  0.18.

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generated by the program for each fit. When curves are properly fit on the X-axis, residuals are expected to be randomly distributed throughout the variable with no drift, i.e., no obvious pattern or long runs of positive or negative values. It is also best for the residuals to have small relative magnitudes. For AarA, the Hill equation was chosen as a preferred model for all three studied rhomboids, suggesting cooperative substrate binding behavior. 10. Calculate the catalytic parameters. Although a discontinuous assay just described can be adapted for the measurements of kinetics, a continuous assay with optical readout detailed in the next section is, in principle, much more powerful.

3.2 Fluorescent Substrates Enzyme substrates that release fluorescent products are commonly used to measure the activity of proteases (Huang, 1991). They offer many advantages starting from high sensitivity and ability to precisely detect even minimal amounts of generated product to the possibility to monitor the reaction in real time and observe the features of enzymatic reaction, including bursts and lag phases. Several fluorogenic substrates were reported and used for rhomboid activity measurements. The Urban group (Dickey et al., 2013) labeled the peptide (1–33 residues) derived from TatA of P. stuartii with FITC at its N-terminus. The kinetic assay for rhomboid using this substrate was performed in proteoliposomes, where both rhomboid protease and TatA-FITC were reconstituted. Unexpectedly, while fluorescence of the substrate was quenched in proteoliposomes, the cleavage of the substrate and release of the fluorophore into the bulk solution restored fluorescence. This enabled detailed kinetic analysis of rhomboid proteolysis within the membrane (Dickey et al., 2013). The distinctive property of rhomboids is that even though their natural substrates are full-length TMDs, the cleavage event relies only on a short recognition motif (Strisovsky et al., 2009). Based on this finding, a fluorogenic substrate consisting of a short peptide (KSp21) derived from substrate of a Drosophila rhomboid Gurken, and FRET donor–acceptor pair, Chromis-645/QSY21 (Pierrat et al., 2011), was developed. The design of the FRET assay is based on the mechanism of energy transfer between two chromophores; from a higher-energy donor to a lower-energy acceptor (F€ orster, 1948). The efficacy of this transfer is inversely proportional to the

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sixth power of the distance between donor and acceptor. If two fluorophores are in close proximity (1–10 nm), the acceptor emission increases because of intramolecular FRET from the donor to acceptor. The time course of KSp21 substrate cleavage by AarA, a rhomboid from P. stuartii showed exponential fit with a linear portion of the curve, from where initial velocity could have been determined. KSp21-based fluorescent assay provided an effective tool for high-throughput screen for rhomboid inhibitors and activators. However, the limited solubility of this substrate impeded the determination of catalytic parameters of its AarA-mediated cleavage (Pierrat et al., 2011). The development of genetically encoded biosensors, such as GFP and related blue, cyan, yellow, and red proteins have overcome such issues as site-specific attachments and solubility of the substrate (Ibraheem & Campbell, 2010). These proteins form FRET pairs with each other or with conventional organic dyes and can be genetically attached to the substrate to form chimera proteins and still retain their fluorescent capability. 3.2.1 Generic Protease Substrate—BODIPY-Casein One of the first substrates used for detection of activity of rhomboid proteases was fluorescently labeled casein. The ability of rhomboids to cleave water-soluble substrates as casein is an apparent paradox, which can be explained by the unusual properties of casein (Wang et al., 2006; Xue & Ha, 2012). It contains a high number of noninteracting proline residues. There are also no disulfide bridges. Therefore, casein has little tertiary structure and it is relatively hydrophobic. It is thought that rhomboids favor loosely packed or even unfolded substrates (Erez & Bibi, 2009; Urban & Freeman, 2003); therefore, these casein features may contribute to its cleavability by rhomboid proteases. Even though casein is not a native substrate it provides a tool to quickly analyze activity (apparent kinetic parameters) for rhomboid proteases (Lazareno-Saez et al., 2013), whose native substrates are not known. We used commercially available Molecular Probes’ EnzChek® Protease Assay Kit (Thermo Fisher Scientific, USA), which contains a casein derivative heavily labeled with the pH-insensitive fluorescent BODIPY dye. Heavy labeling results in almost total quenching of the conjugate’s fluorescence, but once casein is cleaved highly fluorescent BODIPY dye is released. The increase in fluorescence, which can be measured by a fluorimeter, is proportional to protease activity. This approach provides fast and simple technique to monitor the activity in a real time. In our laboratory, we

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developed a detailed kinetic assay using this substrate for assessing catalytic parameters of GlpG, a rhomboid from E. coli (Lazareno-Saez et al., 2013). GlpG is the most characterized rhomboid from structural aspect, but its native substrate repertoire has not been identified. As for any kinetic assay, the initial parameters, including the concentration of enzyme used in the assay, the optimal pH, and detergent concentration, as well as time of reaction should be optimized prior to a kinetics analysis experiment is undertaken. 3.2.1.1 Determination of Optimal Rhomboid Concentration for Kinetic Analysis

Initial rate of the reaction is linearly proportional to the concentration of the enzyme and hyperbolically dependent on substrate concentration. Thus, at high substrate concentrations (approaching saturation), initial reaction rate is only marginally sensitive to substrate concentration. These are suitable conditions for routine assays. 1. Determine the initial concentration of a stock solution of rhomboid protease, using a Pierce BCA protein assay kit (Thermo Fisher Scientific, USA). 2. Make serial dilutions of rhomboid protease using activity buffer (50 mM MES, pH 6.0, 150 mM NaCl, 20% glycerol, 0.1% DDM), where the final enzyme concentrations in the reaction mixture is 0.05, 0.2, 0.5, and 1.5 μM. 3. Prepare a series of activity reaction mixtures in 96-Well Solid Black Corning™ microplate. Each reaction mixture of a final volume of 100 μL should contain the substrate, fluorescently labeled casein (FLcasein), at a fixed final concentration of 10 μM and activity buffer. Preincubate the substrate with the activity buffer at 37°C for 30 min in the dark, because the dye is light sensitive. As we observed, preincubation time is necessary for the system to reach equilibrium, otherwise the assay shows lag phase. 4. Start the series of reactions with the equal volume of rhomboid dilutions. 5. The reaction mixture containing the same concentration of substrate (10 μM) with activity buffer but without enzyme addition treated the same way as sample reactions is used as negative control. This negative control allows subtracting all fluctuations in fluorescence during the time of reaction, which occurs due to the reasons other than the cleavage reaction. 6. Measure the fluorescence emission at 513 nm at 37°C every 5 min during 5 h in fluorescence microplate reader with an excitation wavelength of 503 nm.

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Fig. 5 The time course of FL-casein cleavage by different concentrations of GlpG.

Subtract fluorescence of substrate only from fluorescence of enzymatic reaction at each time point and conduct the relative fluorescent units (RFU) vs time plot. Fig. 5 demonstrates time course of FL-casein cleavage by different concentrations of GlpG. In case of 0.05 μM of GlpG, it is impossible to determine vo, because the concentration of enzyme is too low. In case of 1.5 μM, reaction rate is too high, which impedes vo determination as well. Optimal velocity is observed at 0.2 μM GlpG and this concentration is thus suitable for the subsequent kinetic assays. 3.2.1.2 Determination of Optimal pH for GlpG-Mediated Cleavage of FL-Casein

Several factors important for enzymatic reactions are influenced by pH: substrate binding, its ionization, and the ionization states of the amino acid residues involved in the catalytic reaction. Therefore, determination of pH optimum is a prerequisite for any kinetic assay. Initial rates of FL-casein cleavage by GlpG at broad range of pH were determined. Britton–Robinson universal buffer (Britton & Robinson, 1931) was used to avoid the effect of buffer components on rhomboid activity. Universal buffer consisted of

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mixture of 0.04 M of H3BO3, 0.04 M H3PO4, and 0.04 M CH3COOH, which were adjusted to the desired pH with 0.2 M NaOH. We modified the universal buffer for rhomboids by adding 20% glycerol and 0.1% DDM. 1. Prepare the buffers in pH range between 2 and 11. 2. Prepare a series of activity reaction mixtures in 96-Well Solid Black Corning™ microplate. Each reaction mixture of a final volume of 100 μL contains the substrate, FL-casein, at a fixed final concentration of 10 μM, and a universal buffer with a certain pH. Preincubate the substrate with the universal buffer at 37°C for 30 min. 3. Start the reactions with 0.2 μM (final concentration) of GlpG enzyme. The reaction mixtures containing the same concentration of substrate (10 μM) and buffers with the same pHs as for samples, preincubated for the same period of time at 37°C but without enzyme addition are used as negative controls. 4. Measure the fluorescence emission at 513 nm at 37°C every 5 min during 2 h in fluorescence microplate reader with an excitation wavelength of 503 nm. 5. Subtract fluorescence of substrate only from fluorescence of enzymatic reaction and conduct RFU vs time-dependence plot for each pH value. Determine vo for each curve, using exponential or polynomial fit and estimating the slope of a curve. 6. Plot vo for each pH value vs pH. As it is demonstrated in Fig. 6, pH optimum for GlpG cleavage of FL-casein was at pH 6.0, which is comparable with pH optimum for AarA (pH optimum of which was 6.0 as well). 3.2.1.3 Determination of Optimal Detergent Concentration

Detergent is required in activity buffer at a concentration above its CMC to maintain the solubility of rhomboid proteases. However, the precise detergent concentration for a specific assay is defined by other factors including the nature of substrate, which is a protein itself and also can be influenced by the presence of detergent. Therefore, it is important to assess the optimal detergent concentration for a specific assay (Schroter, Trankle, & Mohr, 2000). We tested the influence of several concentrations of the detergent DDM ranging from 0.05% to 0.75% on the ability of GlpG to cleave FL-casein. 1. Prepare activity reaction mixtures in 96-Well Solid Black Corning™ microplate. Each reaction mixture of a final volume of 100 μL contains the substrate, FL-casein, at a fixed final concentration of 10 μM and activity buffer containing 0.05%, 0.1%, 0.25%, 0.5%, and 0.75% DDM. Preincubate the substrate with the activity buffer at 37°C for 30 min.

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Fig. 6 GlpG activity dependence on pH. The proteolysis reaction was performed at 37°C for 2 h in the presence of 0.2 μM of rhomboid protease and 10 μM of FL-casein in Britton–Robinson universal buffer, at pH interval spanning from 2 to 9.

2. Start the reactions by adding 0.2 μM (final concentration) of rhomboid enzyme. 3. The reaction mixtures containing the same concentration of substrate (10 μM) and activity buffer with the same detergent concentrations as in the reaction samples preincubated for the same period of time at 37°C but without enzyme addition are used as negative controls. 4. Measure the fluorescence emission at 513 nm at 37°C every 5 min during 2 h in fluorescence microplate reader with an excitation wavelength of 503 nm. 5. Subtract fluorescence of substrate only from fluorescence of enzymatic reactions at each time point and conduct RFU vs time-dependence plot for each detergent concentration. Determine vo for each curve, using exponential or polynomial fit and estimating the slope of a curve. As demonstrated in Fig. 7, at 0.1% DDM is the optimal detergent concentration for kinetic assay, with FL-casein as substrate.

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Fig. 7 The dependence of GlpG activity on detergent (DDM) concentration.

3.2.1.4 The Effect of Endogenously Bound Lipids

In the lipid milieu, membrane proteins exist as protein–lipid complexes with specific lipids bound to them. These defined lipid molecules are involved in maintaining structural stability, oligomerization, or assembly of multisubunit complexes and retaining activity of membrane proteins (Dowhan, 1997; Hunte, 2005; Tate, 2001). Endogenous phospholipids copurifying with GlpG are also important for protein stability and activity. It has been shown that each chromatography step can remove up to 50% of endogenous phospholipids from some membrane proteins causing precipitation out of solution (Lemieux, Reithmeier, & Wang, 2002). The effect of delipidation due to SEC on GlpG activity was tested to estimate the optimal conditions for rhomboid kinetic assay. 1. Prepare activity reaction mixtures in 96-Well Solid Black Corning™ microplate. Each reaction mixture of a final volume of 100 μL should contain the substrate, FL-casein, at a fixed final concentration of 10 μM, and activity buffer. Preincubate the substrate with the activity buffer at 37°C for 30 min. 2. Start the reactions by adding 0.2 μM (final concentration) of rhomboid enzyme. Two GlpG samples with equal concentrations were used for this experiment—the protein sample that has been collected after Ni2+-affinity chromatography and tag removal step and a second one, which after the preceding steps was passed through the Superdex 200 13/30 gel filtration column (GE Healthcare Life Sciences, USA).

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3. Use the reaction mixture containing the same concentration of substrate (10 μM) and activity buffer preincubated for the same period of time at 37°C but without enzyme addition as negative control. 4. Measure the fluorescence emission at 513 nm at 37°C every 5 min during 2 h in fluorescence microplate reader with an excitation wavelength of 503 nm. 5. Subtract fluorescence of substrate only from fluorescence of enzymatic reactions at each time point and conduct RFU vs time-dependence plots for GlpG activity prior and after SEC. Determine vo for each curve, using exponential or polynomial fit and estimating the slope of a curve. As demonstrated in Fig. 8, GlpG activity is affected by SEC step. Therefore, it is important to take into consideration the enzyme purification steps when comparing activity of different samples. 3.2.1.5 The Proportionality Between the Fluorescence of FL-Casein and Its Concentration

1. Prepare activity reaction mixtures in 96-Well Solid Black Corning™ microplate. Each reaction mixture of a final volume of 100 μL should

Fig. 8 Activity of GlpG before and after SEC. The vo of GlpG before gel filtration is reflected as 100% activity. The percent of activity is also represented for GlpG after gel filtration.

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contain the FL-casein at a concentration range from 0.15 to 7 μM and activity buffer. Preincubate the substrate with the activity buffer at 37°C for 30 min. 2. Measure the fluorescence emission at 513 nm at 37°C every 5 min during 2 h in fluorescence microplate reader with an excitation wavelength of 503 nm. Conduct RFU vs FL-casein concentration-dependence plot. Fig. 9 demonstrates a linear proportionality between fluorescent signals emitted and the amount of FL-casein used in the kinetic assay, confirming the proper range of substrate concentrations. 3.2.1.6 Can GlpG-Mediated FL-Casein Cleavage Be Fit With Simple Michaelis–Menten Kinetics?

FL-casein is an artificial substrate with no specificity for rhomboid protease. Therefore, it is important to ensure that there is only one major cleavage site for it and FL-casein cleavage by GlpG is a first-order or zero-order reaction. Only in this case, the saturation curve can be fit with Michaelis–Menten equation. The Michaelis–Menten kinetics has both a first-order region, where the rate depends linearly on the substrate concentration and a

Fig. 9 The proportionality between fluorescent signals of different concentrations of FL-casein and the amount of FL-casein used in the kinetic assay are examined.

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zero-order region, when the substrate concentration is high with respect to the KM value. 1. Prepare activity reaction mixtures containing 1 or 10 μM of FL-casein in activity buffer. Preincubate the substrate with the activity buffer at 37°C for 30 min. 2. Start the reactions by adding 0.2 μM (final concentration) of GlpG enzyme. 3. Use the reaction mixture containing 10 μM of FL-casein and activity buffer preincubated for the same period of time at 37°C but without enzyme addition as negative control. 4. Incubate the reaction mixtures at 37°C for 2 h. 5. Add the same amount of SDS-sample buffer to the protein sample. 6. Load 20 μL of a sample on 14% SDS-gel and run the gel. 7. Reveal the fluorescent bands using imaging system for fluorescence. In our lab we use ImageQuant LAS 4000 biomolecular system. As demonstrated in Fig. 10, SDS-gel shows only one major product around 12 kDa, confirming that FL-casein can be used as substrate and the data can be fit with Michaelis–Menten equation. 3.2.1.7 The Kinetic Analysis of GlpG-Mediated Cleavage of FL-Casein

To determine the catalytic parameters of GlpG-mediated cleavage of FL-casein, a real-time optimized kinetic assay was performed as following:

Fig. 10 The SDS-PAGE of FL-casein cleavage by GlpG. Lane 1, fluorescent molecular mass marker; lane 2, FL-casein, no enzyme; lanes 3 and 4, 0.2 μM of GlpG was incubated with 0.2 μM (lane 3) or 10 μM (lane 4) of FL-casein for 2 h at 37°C. Nonfluorescent molecular mass markers are represented on the right.

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1. Prepare activity reaction mixtures in 96-Well Solid Black Corning™ microplate. Each reaction mixture of a final volume of 100 μL should contain different concentrations of FL-casein, ranging from 0.15 to 6 μM and activity buffer. 2. Preincubate the substrate with the activity buffer at 37°C for 30 min. Preincubation time is needed for equilibration of the system, otherwise the assay shows lag phase, which impedes the determination of initial rates. 3. Start the reactions by adding 0.2 μM (final concentration) of GlpG. 4. The set of reaction mixtures containing substrate at the same concentrations as for reaction samples with activity buffer preincubated for the same period of time at 37°C but without enzyme addition are used as negative controls. The negative control allows subtracting all changes in fluorescence during 2 h of reaction, which occur due to the reasons other than the cleavage reaction. 5. Measure the fluorescence emission at 513 nm at 37°C every 5 min during 2 h in fluorescence microplate reader with an excitation wavelength of 503 nm. 6. Subtract fluorescence of substrate only from fluorescence of enzymatic reactions at each time point and conduct RFU vs time-dependence plots for each substrate concentration. Determine vo for each curve, using exponential or polynomial fit and estimating the slope of a curve. 7. To convert the fluorescent units of generated product into μM, perform the cleavage reaction under the same conditions (2 h, 37°C), using the highest and the lowest substrate concentrations. Resolve the cleaved product on SDS-PAGE and visualize the bands using imaging system for fluorescence. Calculate the amount of appeared product in μM, knowing the percentage of cleavage from the fluorescent measurements. Calculate the conversion factor, comparing the amount of produced product in μM to the amount of generated fluorescence. X ¼ F=C ðRFU=μM Þ, where X is a conversion factor; F, generated fluorescence; C, concentration of product in μM. 8. Plot vo vs the corresponding substrate concentration. Fit the data with Michaelis–Menten equation using Prism software (GraphPad, USA) (Fig. 11). Calculate the catalytic parameters.

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Fig. 11 Michaelis–Menten plot of GlpG-mediated cleavage of FL-casein (n ¼ 2, mean  SD). The catalytic parameters were calculated from Michaelis–Menten equation: KM ¼ 0.87  0.2 μM, kcat ¼ 0.307  0.027 min–1.

3.2.2 Fluorescent Transmembrane Protein Substrate—FRET-psTatA The common pair of protein fluorophores for FRET is cyan fluorescent protein (CFP) and yellow fluorescent protein (YFP), which are variants of GFP (Nguyen & Daugherty, 2005). The standard construct design involves a CFP–YFP pair, flanking the peptide bearing sequence of protease cleavage site. When the peptide is cleaved, the two fluorophores get separated, causing the change in FRET signal due to increased emission of the donor (Fig. 12A and B). This phenomenon is used to monitor reaction kinetics (Zauner, Berger-Hoffmann, Muller, Hoffmann, & Zuchner, 2011). The successful design of FRET-based substrate requires close proximity of FRET pair. We took advantage of a small size of one of known rhomboid substrates—TatA protein from P. stuartii (psTatA) to develop FRET-based kinetic assay. psTatA is a 97 amino acid membrane protein with one TMD. It is a component of twin arginine transport (Tat) system and native substrate of AarA, a rhomboid from P. stuartii (Clemmer, Sturgill, Veenstra, & Rather, 2006; Stevenson et al., 2007). It has been demonstrated that rhomboid proteases from E. coli and Haemophilus influenzae can also process

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Fig. 12 (A) The schematic representation of FRET-based assay. (B) SDS-PAGE of protein sample fractions after SEC purification of psTatA-FRET. (C) Emission spectrum of psTatAFRET. psTatA-FRET (2 μM) before (black) and after (gray) incubation with the same concentration of AarA rhomboid for 4 h at 37°C.

psTatA in vitro, however, with different rates (Arutyunova et al., 2014). In this chapter, we will describe the development of kinetic assay for catalytic parameters determination of rhomboid-mediated enzymatic reaction for the only known prokaryotic physiological pair of AarA rhomboid–psTatA substrate. FRET-based assays are used to study proteolytic activity in vitro and in vivo (Fields, 2001; Knight, 1995; Lombard, Saulnier, & Wallach, 2005), but such assays are not common for intramembrane proteases. Often the ratio of acceptor vs donor emission is used as a quantitative parameter to monitor and determine protease activity. However, such ratiometric analysis does not directly correlate with the amount of digested substrate due to signal cross-contaminations at both acceptor and donor emission wavelengths by acceptor and donor self-fluorescence. We use a modified method for FRET data analysis to avoid signal cross-contamination and accurately

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determine kinetic parameters of substrate digestion (Liu, Song, Madahar, & Liao, 2012). Absolute fluorescence signals contributed by the donor and acceptor, and FRET at the acceptor and emission wavelengths were quantified, enabling FRET data to be converted into protein concentration with high precision. The entire protein sequence of psTatA was cloned between engineered FRET pair—CyPet and YPet, which are the derivatives of CFP and YFP (Nguyen & Daugherty, 2005) in pBad-Myc-HisB plasmid. The schematic representation of the construct is shown in Fig. 12A. pBAD-Myc-HisB vector was employed to express the construct. The construct contained 6Histag for detection and purification purposes. 3.2.2.1 Expression and Purification of psTatA-FRET Substrate

1. Transform pBad-Myc-His vector, bearing CyPet-psTatA-YPet construct into K12 GlpG knockout competent cells (glpG::Kn) from Keio library and select transformants on LB plates containing 100 μg/mL of ampicillin. We use E. coli Keio Knockout Collection cells (Baba et al., 2006) in order to eliminate the cleavage of expressed psTatA-FRET substrate by endogenous GlpG. Even though psTatA is not a native substrate for E. coli rhomboid, it has been demonstrated that GlpG is able to cleave psTatA (Strisovsky et al., 2009). 2. Pick a single transformant colony and inoculate into 150 mL of LB medium, supplemented with 100 μg/mL of ampicillin (LB-Amp). Shake culture overnight (200 rpm) at 37°C. The following day subinoculate 20 mL of the overnight culture into six individual 4 L flasks containing 1 L of LB-Amp. 3. Grow cells at 37°C until OD600 reaches 0.6, induce the protein with 0.02% arabinose, and express at 24°C for overnight. 4. Harvest the cells, resuspend the cell pellet in 1:4 (weight:volume) ratio of 50 mM Tris–HCl, pH 8.0, 200 mM NaCl buffer, containing Protease Inhibitor tablet(s) and 1 μg/mL (final concentration) of DNaseI, and lyse the cells. 5. Centrifuge the lysed cell extract at 20,000  g for 30 min at 4°C to remove unbroken cells and cell debris. Add DDM detergent to the supernatant to the final concentration of 0.1% and incubate with gentle stirring for 30 min at 4°C. Although psTatA has TMD and if expressed alone it is purified from the membrane fraction (see Section 3.2.2.1), when fused to soluble CyPet and YPet protein, it is found mostly in the supernatant. However, the presence of highly hydrophobic region

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of membrane domain in aqueous solution is energetically unfavorable and can cause protein aggregation. In order to mask the protein’s hydrophobic regions, the detergent is added. For rhomboids, DDM is used for solubilization and purification; therefore, we preferred to use the same detergent for the substrate to keep the kinetic assay components uniform. Add the supernatant with detergent to Ni2+-resin preequilibrated with IMAC buffer (50 mM Tris–HCl, pH 8.0, 200 mM NaCl, 20% glycerol, 0.1% DDM). Incubate the sample at 4°C on a shaker at low speed for 2 h. The batch method of sample application maximizes the contact time of fusion protein with the resin for the most efficient binding. The amount of required resin should be determined empirically. We use 0.5 mL of settled Ni2+-resin per each liter of cell culture with expressed psTatA-FRET. Chelating agents such as EDTA or citrate should not be included in any buffer. Load sample/resin mix onto the 25-mL drip column, collect flowthrough for analysis and wash the resin with 20 CV (column volume) of wash buffer (50 mM Tris–HCl, pH 8.0, 500 mM NaCl, 20% glycerol, 30 mM imidazole, 0.1% DDM). Elute the protein fractions in a stepwise manner with 2–5 CV of IMAC buffer containing 100, 250, 500, 1000, and 2000 mM imidazole, respectively. Perform SDS-PAGE analysis of eluted fractions and combine the fractions according to purity and amount of protein of interest. Note that the fusion to FRET pair allows fluorescent detection of psTatA-FRET substrate directly on SDS-PAGE. It permits the rapid monitoring of total protein patterns during the whole purification process. The only requirement for sample preparation for SDS-PAGE is using unboiled samples in sample buffer. Concentrate the combined protein sample using 30 kDa MWCO filtration device (EMD Millipore, USA) to 1–3% of SEC column that is expected to be used for the next step. Note that the molecular weight of DDM micelles is around 50 kDa, therefore, the detergent concentration increases during protein concentration. Therefore, it is very important to avoid overconcentration to prevent protein aggregation and denaturation due to high detergent concentration. Load concentrated protein sample on gel filtration column, Superdex 200 16/60 (GE Healthcare Life Sciences, USA), equilibrated with 20 mM Tris–HCl, pH 8.0, 200 mM NaCl, 5% glycerol, 1 mM EDTA, 0.1% DDM. Collect protein fractions. Analyze them on SDS-PAGE

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and combine the most pure ones. Concentrate to appropriate concentration using 30 kDa MWCO filtration device (EMD Millipore, USA). Assess the protein concentration with the (BCA) protein assay kit (Thermo Fisher Scientific, USA). As demonstrated in Fig. 12, B the substrate fractions after SEC are relatively pure, which is very important for future kinetic assay. 11. Aliquot and flash-freeze the protein samples. Store them at –80°C. 3.2.2.2 The Reliability of psTatA-FRET for FRET-Based Kinetic Assay

To test if the purified substrate can be used for FRET-based analysis, we took an emission spectrum before and after psTatA-FRET cleavage by AarA rhomboid. 1. Prepare 2 μM solution of psTatA-FRET in activity buffer (50 mM MES, pH 6.0, 150 mM NaCl, 20% glycerol, 0.1% DDM). 2. Start the enzymatic reaction by addition of the same amount (2 μM) of AarA rhomboid. Incubate the reaction mixture at 37°C for 4 h. 3. Read the emission spectrum of 2 μM psTatA-FRET alone (Fig. 12C, black) and psTatA-FRET after incubation with AarA (Fig. 12C, gray). A significant change in fluorescent signal was observed between the undigested and digested substrate. Two peaks can be seen for uncleaved CyPet-psTatA-YPet substrate showing partially quenched fluorescence of CyPet at 475 nm and full fluorescence of YPet at 530 nm. Upon substrate cleavage, the fluorescence emission of CyPet at 475 nm is increased, whereas the YPet emission at 530 nm is decreased due to the loss of FRET energy transfer. This result showed that psTatA-FRET is a reliable substrate for determination of kinetic parameters of AarA. 3.2.2.3 Optimization of FRET-Based Kinetic Assay Parameters

The objective of any kinetic assay is to measure the maximal enzyme activity under defined conditions, so that the activity can be compared between different samples or even between different laboratories. Even though in vitro conditions often do not closely resemble those in the cell, the determination and optimization of all parameters affecting enzyme’s activity aids in understanding the activity occurring in vivo. These parameters include pH, temperature, substrate concentration(s), and ionic strength and nature of compounds present in activity buffer. Activity is measured as the initial rate of substrate consumption when no product is present and optimization of proper conditions for initial rate determination is the prerequisite of any successful kinetic analysis. It is

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impossible to predict the concentration of enzyme required for obtaining the maximal initial rate and it should be done empirically. 3.2.2.4 Determination of Optimal Rhomboid Concentration for FRET-Based Kinetic Analysis

Based on Michaelis–Menten equation (1) it is clear that the main factor that determines the enzymatic reaction is the substrate concentration [S]. vo ¼ Vmax ½S=½S + KM :

(1)

The initial rate of reaction is proportional to enzyme’s concentration, therefore it is crucial to define the conditions for optimal enzyme’s concentration determination. The simplest approach is to use high substrate concentration so it practically does not change during the reaction time: ½So   Δ½St   ½So ,

(2)

where [So] is the initial substrate’s concentration, Δ[St] is the change in substrate concentration during reaction time, t. If the substrate concentration is too low ([S]  KM), then the reaction rate decreases impeding initial rate determination. 1. Determine the initial concentration of a stock solution of rhomboid protease, using Pierce BCA protein assay kit (Thermo Fisher Scientific, USA). 2. Make a serial dilutions of rhomboid protease using activity buffer (50 mM MES, pH 6.0, 150 mM NaCl, 20% glycerol, 0.1% DDM) so the final enzyme concentrations in the reaction mixture is 0.05, 0.15, 0.3, 0.5, and 1.5 μM. 3. Prepare a series of activity reaction mixtures in 96-Well Solid Black Corning™ microplate. Each reaction mixture of a final volume of 100 μL should contain the substrate, psTatA-FRET, at a fixed final concentration of 7 μM, and activity buffer. Preincubate the substrate with the activity buffer at 37°C for 30 min. As we observed, preincubation time is necessary for the system to reach equilibrium, otherwise the assay shows lag phase. Indeed, the term initial rate is referred to the steady-state rate of the reaction, which is established after any “presteady-state” events have occurred. 4. Start the series of reactions with the equal volume of rhomboid protease with different concentrations. 5. The reaction mixture containing the same concentration of substrate (7 μM) with activity buffer, preincubated for the same period of time at 37°C but without enzyme addition is used as negative control.

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This negative control allows subtracting all changes in fluorescence during 2 h of reaction, which occur due to the reasons other than the cleavage reaction. 6. Measure the emission intensity of CyPet at 475 nm and YPet at 530 nm with the excitation wavelength of 414 nm for enzymatic reactions and for negative control sample in a fluorescence multiwell plate reader (we use SynergyMx, BioTek) each 5 min for 5 h. 7. Subtract fluorescence of substrate only from fluorescence of enzymatic reaction at each time point and conduct the RFU vs time plot. Fig. 13, A demonstrates time course of psTatA-FRET cleavage by different concentrations of AarA. In case of 0.05 μM of AarA it is impossible to determine vo, because the concentration of enzyme is too little.

Fig. 13 (A) The time course of psTatA-FRET cleavage by different concentrations of AarA. (B) The dependence of initial velocity of psTatA-FRET cleavage performed by different concentration of AarA on substrate concentration.

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In case of 1.5 μM, reaction rate is too high, which impedes vo determination as well. 0.15 and 0.3 μM of enzyme give the optimal time course curve—the linear part lasts for at least 120 min and the velocity reaches the maximum within 180 min. If the initial velocity determination experiment is conducted under optimal assay conditions, the dependence between the enzyme concentration and vo is linear, as it is shown in Fig. 13B. Thus, rhomboid concentration suitable for kinetic analysis was determined to be in the range between 0.15 and 0.3 μM. 3.2.2.5 Dependence of Rhomboid Activity on pH

pH has a strong effect on enzyme’s activity by changing the ionization state of acidic or basic amino acids. This can lead to alteration of ionic bonds, which determines overall 3 days protein structure and subsequently to loss of substrate recognition and/or activity. pH also can affect the substrate’s properties. It is especially important for proteases, which substrates are proteins. The pH where the enzyme is the most active is known as optimum pH. 1. In order to find the optimum pH, vo is determined at broad range of pH maintaining all other assay parameters constant and optimal. To avoid the effect of buffer components on rhomboid activity we chose to work with Britton–Robinson universal buffer (Britton & Robinson, 1931). Universal buffer consists of mixture of 0.04 M of H3BO3, 0.04 M H3PO4, and 0.04 M CH3COOH, which are adjusted to the desired pH with 0.2 M NaOH. We modified the universal buffer for rhomboids by adding 20% glycerol and 0.1% DDM to it. 2. Prepare the buffers with pH range between 2 and 11. 3. Prepare a series of activity reaction mixtures in 96-Well Solid Black Corning™ microplate. Each reaction mixture of a final volume of 100 μL should contain the substrate, psTatA-FRET, at a fixed final concentration of 7 μM and a universal buffer with a certain pH. Preincubate the substrate with the universal buffer at 37°C for 30 min. 4. Start the reactions with 0.2 μM (final concentration) of AarA enzyme. 5. The reaction mixtures containing the same concentration of substrate (7 μM) and buffers with the same pHs as for samples, preincubated for the same period of time at 37°C but without enzyme addition are used as negative controls. 6. Measure the emission intensity of CyPet at 475 nm and YPet at 530 nm with the excitation wavelength of 414 nm for enzymatic reactions and for negative control samples in a fluorescence multiwell plate reader (we use SynergyMx, BioTek, Winooski, USA) each 5 min for 2 h.

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7. Subtract fluorescence of substrate only from fluorescence of enzymatic reaction at each time point and conduct RFU vs time-dependence plot for each pH value. Determine vo for each curve, using exponential or polynomial fit and estimating the slope of a curve. 8. Plot vo for each pH value vs pH. As it is demonstrated in Fig. 14, pH maximum for AarA rhomboid was determined to be at pH 6.0. 3.2.2.6 Dependence of Rhomboid Activity on Temperature

Enzymatic reactions, like most chemical reactions, are temperature dependent. As temperature increases, reacting molecules have more kinetic energy; this increases the chance of collision and accelerates the rate of reaction. There is a certain temperature at which an enzyme’s catalytic activity is at its greatest. Above this temperature enzyme’s structure begins to collapse and even small changes at the remote from active-site regions could cause change in activity due to altered substrate–enzyme initial interaction and recognition. The effect of temperature on rhomboid’s activity with different substrates is well described in the recent paper from our laboratory (Panigrahi et al., 2016).

Fig. 14 AarA activity dependence on pH. The proteolysis reaction was performed at 37° C for 2 h in the presence of 0.18 μM of rhomboid protease and 2 μM of psTatA-FRET in Britton–Robinson universal buffer, at pH interval spanning from 2 to 9.

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To determine the optimal temperature for rhomboid’s kinetic assay three temperatures were tested: 24°C, 30°C, and 37°C. 1. Prepare activity reaction mixtures in 96-Well Solid Black Corning™ microplate. Each reaction mixture of a final volume of 100 μL should contain the substrate, psTatA-FRET, at a fixed final concentration of 7 μM, and activity buffer. Preincubate the substrate with the activity buffer at the temperature of reaction for 30 min. 2. Start the reactions by adding 0.2 μM (final concentration) of AarA enzyme. 3. The reaction mixtures containing the same concentration of substrate (7 μM) with activity buffer, preincubated for the same period of time at 24°C, 30°C, or 37°C but without enzyme addition are used as negative controls. 4. Measure the emission intensity of CyPet at 475 nm and YPet at 530 nm with the excitation wavelength of 414 nm for enzymatic reactions and for negative control samples in a fluorescence multiwell plate reader (we use SynergyMx, BioTek) each 5 min for 2 h at 24°C, 30°C, and 37°C. 5. Subtract fluorescence of substrate only from fluorescence of enzymatic reaction at each time point and conduct RFU vs time-dependence plot for each reaction temperature. Determine vo for each curve, using exponential or polynomial fit and estimating the slope of a curve. As it is demonstrated in Fig. 15, the optimal temperature for AarA-mediated cleavage of psTatA is 37°C, lower temperatures give decreased initial velocity. Higher temperatures were not tested for optimization of activity assay conditions because it was outside of physiological range for this enzyme.

3.2.2.7 Determination of Optimal Detergent Concentration

The concentrations of detergent could affect the enzymatic activity of membrane proteins, the substrate stability, or the specific signal in concentration response curve (Schroter et al., 2000). Therefore, the optimal concentration of detergent may not be the same for different substrates. We tested the influence of several DDM concentrations ranging from 0.05% to 0.75% on AarA activity with psTatA-FRET substrate. 1. Prepare activity reaction mixtures in 96-Well Solid Black Corning™ microplate. Each reaction mixture of a final volume of 100 μL should

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Fig. 15 The time course of AarA-mediated cleavage of psTatA-FRET at different temperatures. 0.2 μM of AarA was incubated with psTatA-FRET, at a final concentration of 7 μM in activity buffer for 2 h at 24°C, 30°C, and 37°C.

2. 3.

4.

5.

contain the substrate, psTatA-FRET, at a fixed final concentration of 7 μM and activity buffer containing 0.05%, 0.075%, 0.1%, 0.25%, 0.5%, and 0.75% DDM. Preincubate the substrate with the activity buffer at 37°C for 30 min. Start the reactions by adding 0.2 μM (final concentration) of rhomboid enzyme. The reaction mixtures containing the same concentration of substrate (7 μM) and activity buffer with the same detergent concentrations as in the reaction samples, preincubated for the same period of time at 37°C but without enzyme addition are used as negative controls. Measure the emission intensity of CyPet at 475 nm and YPet at 530 nm with the excitation wavelength of 414 nm for enzymatic reactions and for negative control samples in a fluorescence multiwell plate reader (we use SynergyMx, BioTek) each 5 min for 2 h at 37°C. Subtract fluorescence of substrate only from fluorescence of enzymatic reaction at each time point and conduct RFU vs time-dependence plot for detergent concentration. Determine vo for each curve, using exponential or polynomial fit and estimating the slope of a curve. As demonstrated in Fig. 16, at 0.1% DDM AarA-mediated cleavage of psTatA is at its highest rate, whereas higher detergent concentrations inhibit the enzymatic reaction.

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Fig. 16 AarA activity dependence on DDM concentration. 0.2 μM of AarA was incubated with psTatA-FRET, at a final concentration of 7 μM in activity buffer, having different concentrations of DDM for 2 h at 37°C.

3.2.2.8 The Proportionality Between the Fluorescence Emitted and the Substrate Concentration

FRET-based substrates is a convenient and sensitive approach to study the kinetics of proteases; however, the main drawback of these assays is that at increasing concentrations of substrate (usually higher than 20 μM) its increasing absorbance introduces the inner filter effect that decreases the fluorescence emission and change (Palmier & Van Doren, 2007). Consequently, the dependence of FRET-substrate fluorescence on its concentration becomes a problem, which leads to understated kinetic parameters. To ensure that fluorescence change does not lose linearity at substrate concentrations used in our assay, we determined the proportionality between fluorescent signal and the amount of psTatA-FRET. 1. Prepare activity reaction mixtures in 96-Well Solid Black Corning™ microplate. Each reaction mixture of a final volume of 100 μL should contain the substrate, psTatA-FRET, at a concentration range from 0.13 to 7 μM, and activity buffer. Preincubate the substrate with the activity buffer at 37°C for 30 min. 2. Measure the emission intensity of CyPet at 475 nm with the excitation wavelength of 414 nm in a fluorescence multiwell plate reader at 37°C. 3. Conduct RFU vs psTatA-FRET concentration-dependence plot. Fig. 17 demonstrates a linear proportionality between fluorescent signals

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emitted and the amount of psTatA-FRET used in the kinetic assay, confirming the proper range of substrate concentrations. 3.2.2.9 Determination of Catalytic Parameters of AarA-Mediated Cleavage of psTatA-FRET

To study the AarA-mediated cleavage of psTatA in detail, a real-time optimized FRET-based kinetic assay was utilized. For the FRET data analysis, we used a modified method, which allows an accurate determination of catalytic parameters taking into account autofluorescence to avoid signal crosscontamination (Liu et al., 2012). 1. Prepare activity reaction mixtures in 96-Well Solid Black Corning™ microplate. Each reaction mixture of a final volume of 100 μL should contain different concentrations of substrate, psTatA-FRET, ranging from 0.13 to 7 μM and activity buffer. 2. Preincubate the substrate with the activity buffer at 37°C for 30 min. Preincubation time is needed for equilibration of the system, otherwise the assay shows lag phase, which impedes the determination of initial rates, because initial rate is referred to the steady-state rate of the reaction, which is established after any “presteady-state” events. 3. Start the reactions by adding 0.2 μM (final concentration) of AarA enzyme. 4. The set of reaction mixtures containing substrate at the same concentrations as for reaction samples with activity buffer preincubated for the same period of time at 37°C, but without enzyme addition are used as negative controls. The negative control allows subtracting all changes in fluorescence during 2 h of reaction, which occur due to the reasons other than the cleavage reaction.

Fig. 17 Proportionality between psTatA-FRET fluorescence and concentration.

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5. Measure the emission intensity of CyPet at 475 nm and YPet at 530 nm with the excitation wavelength of 414 nm in a fluorescence multiwell plate reader (SynergyMx, BioTek) every 5 min for 2 h at 37°C. 6. To negate the effect of signal cross-contamination, determine the CyPet and YPet direct emissions and total emissions at 530 nm. The emission of the recombinant protein CyPet-psTatA-YPet is measured at 475 nm when excited at 414 nm to determine the CyPet direct emission; the emission is measured at 530 nm when excited at 475 nm to determine the YPet direct emission. 7. Conduct RFU vs time plot for each substrate concentration by subtracting fluorescence of substrate only from fluorescence of enzymatic reaction at each time point. 8. Obtain digested concentration of substrate using the following equation, developed by Liu et al. (2012):   M x 0     FL530  αFLCyPet 475  βFLYPet 530 FL530 ¼ 414 M 414 475 414   30 + α kðM  xÞ + jx + βFLYPet530 68 475 FL530 and FL0530 are total fluorescence emission at 530 nm when 414 414

excited at 414 nm before and after digestion, respectively, M is the total amount of CyPet-psTatA-YPet, and x is the amount of digested CyPet-psTatA-YPet. α is ratio of fluorescence emission by CyPet at 530–475 nm under excitation at 414 nm, whereas αFLCyPet475 is 414

CyPet direct emission at 475 nm when excited at 414 nm. Similarly β is ratio of fluorescence emission by YPet at 530–475 nm under excitation at 475 nm. Calculate YPet direct emission (βFLYPet530 ) at 530 nm when 475

excited at 475 nm using this ratio. 30/68 gives the molecular mass ratio of CyPet-psTatA to CyPet-psTatA-YPet. The plot of emission of CyPet-TatA-YPet at 475 nm under excitation at 414 nm vs amount yields a straight line; the constant k determines the slope of this line. Similarly, there is a linear relationship between the emission of CyPetpsTatA + YPet (1:1 M ratio) under excitation at 414 nm and the protein amount is found. In this case, j describes the slope of the plot. Both parameters, j and k, were calculated from standard plots of fluorescence emission vs amount of protein, and used in the equation.

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9. Using converted values conduct the plot of product increase vs time for each substrate concentration. Determine vo for each curve, using exponential or polynomial fit and estimating the slope of a curve. 10. Plot vo vs the substrate concentration. For AarA, we fit the plot with both Michaelis–Menten and the Hill equations using Prism software, GraphPad, USA (Fig. 18). Choose the preferred fit based on R2 values and analysis of the residuals; these parameters are generated by the program for each fit. When curves are properly fit, on the X-axis residuals are expected to be randomly distributed throughout the variable with no drift, i.e., no obvious pattern or long runs of positive or negative values. It is also best for the residuals to have small relative magnitudes. For AarA, the Hill equation was chosen as a preferred model for all three studied rhomboids, suggesting cooperative substrate binding behavior. The Hill coefficient (h) indicates the degree of cooperativity. The Hill coefficient for AarA was h ¼ 1.4  0.1. 11. Calculate the catalytic parameters.

Fig. 18 (A) Overlay of Michaelis–Menten and Hill plots for kinetic data of AarA-mediated cleavage of psTatA-FRET. The catalytic parameters derived from Hill equation: K0.5 ¼ 3.9  0.3 μM, kcat ¼ 0.17  0.07 min–1, Hill coef. ¼ 1.8  0.1. (B and C) Residual plots of Hill (B) and Michaelis–Menten (C) equations for kinetic models accuracy validation.

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ACKNOWLEDGMENTS M.J.L. acknowledges support from Canadian Institute for Health Research (CIHR:MOP 93557), Alberta Innovates Health Solutions (AIHS), Parkinson’s Society of Canada, and the Natural Sciences and Engineering Research Council—International Research Training Group in Membrane Biology. Infrastructure used in this work was funded by the Canadian Foundation for Innovation. K.S. acknowledges support from EMBO (Installation Grant no. 2329), Ministry of Education, Youth and Sports of the Czech Republic (projects no. LK11206 and LO1302), Marie Curie Career Integration Grant (project no. 304154), and the National Subvention for Development of Research Organisations (RVO: 61388963) to the Institute of Organic Chemistry and Biochemistry.

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AUTHOR INDEX Note: Page numbers followed by “f ” indicate figures.

A Aach, J., 380–381 Abdiche, Y.N., 81 Abdine, A., 208–225 Abe, H., 65 Abidi, N., 81 Abulrob, A., 85 Ackaert, C., 87–88 Ackerman, E., 159–160 Ackerman, S., 316 Acx, H., 68–69 Adachi, A., 158 Ades, S.E., 2–4 Adrain, C., 256–257, 397–398 Aerts, A., 87–88 Aeschbach, L., 138, 144–145 Aggeler, P.M., 359 Ahmad, I., 27–29 Ahmadvand, D., 80–81 Ahn, K., 175, 258 Ahn, S., 85–86 Akabas, M.H., 189–193 Akazawa, Y., 78 Akiyama, K., 2–32 Akiyama, S., 2–4 Akiyama, Y., 2–32, 120–121, 210–212, 230–231, 256–257, 282–286, 287f, 396–400 Alattia, J.R., 144–145 Alba, B.M., 2–4 Albaghdadi, H., 85 Albeck, A., 280 Alexandrov, A., 214–216 Alguel, Y., 333–334 Allford, S.L., 377–378 Almo, S.C., 340 Almonacid, D.E., 340 Alva, V., 310, 340–341 Am Ende, C.W., 175 Amara, S.G., 189–191 Amelotti, J.M., 353 Amour, A., 49–50 Amstutz, P., 63–64

Andersen, K.K., 86–87 Andreeva, A., 145–146, 248, 257–258, 283f, 285–286, 287f, 396 Andrews, D., 313–314 Angala, S.K., 311–313 Angsuthanasombat, C., 84 Annaert, W., 84, 128–129, 137, 186–187, 197–198, 230–231, 296 Anne, J., 38–39 Anraku, Y., 337 Ansell, K., 286, 287f, 289, 396–398, 408–409 Apfeld, J., 187–188 Appel, A., 102–104, 256–257 Ara, T., 320, 401, 421 Arakawa, T., 65 Arawaka, S., 187–188 Arbabi, M., 85 Arimon, M., 61, 76, 85, 193–194 Arold, N., 120 Arutyunova, E., 100–101, 116, 256–274, 396–433 Arvan, P., 361–362 Ashby, M.N., 316 Asher, S.A., 210, 221–222 Ashman, K., 128–130, 187–188, 230, 297 Ashwell, C.M., 39–41 Assur, Z., 136–137 Atkins, W.M., 65 Attersand, A., 311, 316 Auer, M., 259–260 Augustin, A., 163 Augustin, M., 83 Austin, J.C., 209–210 Auzanneau, F.I., 50–51 Azerad, R., 356–357 Azuma, K., 379

B Baatout, S., 87–88 Baba, M., 320, 401, 421 Baba, T., 320, 401, 421 439

440 Babbitt, P.C., 340 Baches, S., 175 Bachovchin, D.A., 397 Bacskai, B.J., 60–61 Baert, K., 370 Baetselier, P., 87–88 Bai, X., 61, 101–102 Bai, X.-C., 2, 142f, 144–145, 146f, 147–148, 189, 191–194, 208–209, 297 Bai, Y., 315–316, 319, 330, 334, 337 Baker, E.N., 36, 38 Baker, M., 130 Baker, R.P., 8, 100–101, 130–131, 135–136, 145–146, 146f, 211–212, 230–250, 256–258, 280–284, 289–290, 301–305, 396–400, 408 Baker, T.A., 25 Bakkers, J., 313–314 Balakrishnan, G., 210, 223–224 Balasubramanian, S., 144–145 Ball, C.R., 87 Ballard, T.E., 175 Bammens, L., 60–61, 144–145, 188, 196–197, 298 Baneyx, F., 259–260 Bang, A.G., 128 Bangham, A.D., 238–239 Bangphoomi, K., 84 Baranski, T.J., 195–196 Barbosa, M.D., 43, 47–48 Barford, D., 128–129 Barkocy-Gallagher, G.A., 46–47 Barnych, B., 87 Barrett, A.J., 129–130, 280–282 Barretto, R., 380–381 Barzykin, A.V., 233–235 Basset, G., 137–138, 176, 187–188 Bassford, P.J., 46–47 Baszis, S.R., 314 Bateman, K.S., 102–104, 131, 135–136, 145–146, 231, 256–259, 280–282, 396 Bates, M., 84–85 Batot, G., 36, 38 Baty, D., 75 Bauer, C.E., 315 Baumann, K., 159–160, 163, 171, 175–176 Baumeister, R., 163, 167 Bayburt, T.H., 65

Author Index

Beale, S.I., 315 Beckwith, J., 368–369 Began, J., 100–101, 145–146, 281f, 282–285, 283f, 287f, 288–290, 396 Beher, D., 158–159 Beirnaert, E., 86–87 Beis, K., 265–266, 333–334 Bell, R.G., 352–353 Belvo, M.D., 39 Ben Abderrazek, R., 81 Ben-Chetrit, N., 80–81 Benilova, I., 60–61, 144–145, 188, 196–197, 298 Benlasfar, Z., 81 Benoit, E., 354, 370, 376–378 Ben-Shem, A., 131, 135–136, 145–146, 208–209, 231, 257–258 Benurwar, M., 60–61, 68–69, 144–145, 159–160, 188, 196–197, 298 Berezovska, O., 60–61, 159, 175, 189–191, 194–195, 198, 289–290 Berg, D.T., 361 Berg, S., 314, 337 Bergbold, N., 129 Berger, E.P., 186–187 Berger-Hoffmann, R., 419 Berkner, K.L., 359–361 Bever, C.S., 87 Beyer, P., 314, 316 Bhatt, S., 239–240, 282–284, 400, 419–420 Bibi, E., 128, 131, 135–136, 145–146, 208–209, 231, 257–258, 409 Bibl, M., 144–145 Bicknell, R., 285–286 Biegert, A., 316–319 Bilgin, N., 45 Bilic, S., 87 Bineva, G., 128–129 Binns, K., 128–130, 187–188, 230, 297 Biswas, K., 311–313 Black, M.T., 39 Blackburn, M.N., 361–362 Blain, J.F., 159 Blais, D.R., 100–101, 112–114 Blanchetot, C., 87 Bledsoe, P.J., 358–359 Blennow, K., 186–187 Block, H., 107–110

441

Author Index

Blumbach, K., 358–359 Bockenhauer, S., 61 Bockstael, K., 84 Boehm, R., 311 Boerman, O., 87–88 Bogdanove, A.J., 372 Bogue, S., 311–313, 319 Bohm, C., 61, 193–194 B€ ohm, S., 315–316 B€ oldicke, T., 85–86 Bolduc, D.M., 8, 240, 296–307 Boltz-Nitulescu, G., 84 Bond, J.S., 280–282 Bonitz, T., 310, 340–341 Bonnert, T.P., 70 Booher, N.J., 372 Boons, E., 84 Booth, C.R., 148 Booth, P.J., 264–265 Borch, J., 65 Borchelt, D.R., 186–187, 195–196 Borgers, M., 60–61, 144–145, 188, 196–197, 298 Bosch, L., 87 Bosse, R., 78 B€ ottcher, J., 83 Bouchard, B.A., 378–379 Bouloc, N., 286, 287f, 289, 396–398, 408–409 Boutton, C., 86 Bouwens, L., 85 Bowman, S., 313–314 Boyce, B., 350–351 Boyd, D., 368–369 Boyne, M.E., 311–313 Bradley, M., 49–50 Brandenburg, V., 357 Brandt, W., 313–314, 323–324, 334, 337 Branston, S.D., 71 Brase, S., 130 Br€auer, L., 323–324, 334, 337 Braunfeld, M.B., 148 Breddam, K., 50–51 Breiteneder, H., 84 Brendel, M., 171 Brendel, V.P., 372 Brenner, B., 379 Breteler, M.M., 186–187

Brimble, M.A., 36, 38 Britton, H.T.S., 411–412, 426 Brockhaus, M., 163 Bron, S., 36 Bronstein, M., 379 Brooks, C.L., 145–146, 258–259 Broun, A., 340 Brouwer, A.J., 100–101, 285–289, 287f, 396 Brown, C.L., 356–357 Brown, M., 208–225 Brown, M.C., 210 Brown, M.S., 2, 36, 128–130, 208–209, 230, 296 Brown, S.J., 397 Brubaker, G., 39–41 Bruckner, R., 43, 47–48 Brunak, S., 46–47 Bruton, G., 39 Buelens, K., 61–62 Bugelski, P., 361–362 Buldyrev, I., 198, 289–290 Bulic, B., 175 Bullinger, P.R., 43 Bumba, L., 83 Burgess, A.I., 356–357 Burns, M.P., 163 Bursavich, M.G., 159 Bush, M., 101–102 Bussow, K., 138 Butt, T.R., 216–217 Byrne, B., 333–334

C Cahoon, E.B., 314 Cahuzac, N., 130 Cai, T., 189–191, 193–197 Cai, W., 87–88 Callewaert, N., 63–64 Cameron, A.D., 265–266, 328–329, 333–334 Camire, R.M., 360–361 Campbell, R.E., 409 Cantor, A.B., 356–357 Cao, E., 148 Cao, G., 36, 40, 43 Capell, A., 137, 176 Cappello, C.A., 143–145, 301–302 Carlisle, T.L., 356, 360–361

442 Carlone, G.M., 366 Carlos, J.L., 39–41, 45 Carneiro, J.D., 383–386 Carpenter, E.P., 265–266 Casareno, R., 43 Castro, J.L., 128–129, 186–187, 194, 296 Castro, S.V., 175 Cataland, S., 87 Catanzaro, V., 313–314 Caveliers, V., 75, 87–88 Ceccarelli, E.A., 269, 272–274 Chaba, R., 2–4 Chaicumpa, W., 84 Chakravarty, R., 87–88 Chames, P., 75 Chan, H.K., 39–40 Chang, B.I., 285–286 Chang, C.Y., 352, 370 Chang, G., 63–64 Changeux, J.-P., 61 Chao, Y.B., 361 Chatterjee, D., 311–313 Chatterjee, S., 41, 43 Chau, D.M., 175 Cha´vez Gutierrez, L., 60, 62, 159, 194 Chavez, J., 136–137, 208–225 Cha´vez-Gutierrez, L., 60–88, 144–145, 188–191, 196–197, 298 Checler, F., 143 Chen, B.R., 63–64 Chen, F., 135 Chen, G., 2–4 Chen, M.C., 210 Chen, T.F., 80–81 Chen, X., 128–130, 296 Chen, Y., 258–260 Chen, Z., 83–84 Cheng, D., 129–130 Cheng, S., 159–160 Cheng, S.Y., 46–47 Cheng, T.L., 285–286 Cheng, W., 311, 316, 334, 337, 339–340 Cheng, Y., 148 Cheong, J.-J., 314 Cherepanov, P.P., 322 Cherezov, V., 335–337

Author Index

Chernaia, M., 36, 40 Cherney, M.M., 102–104, 131, 135–136, 145–146, 231, 256–259, 280–282, 396 Cherry, S., 272–274 Chevet, E., 78 Chi, N.C.N., 313–314 Chi, Z., 210 Chiba, S., 2–4, 7, 11, 14 Chin, J.K., 39–40 Chin, J.W., 27 Cho, S., 8, 100, 135–136, 145–146, 233–236, 239–240, 244–246, 248–250, 282–285, 287f, 288–290, 301–305, 396–398, 408 Choudhury, H., 256–258 Chow, K.Y., 87 Chowdhury, S., 285–286, 287f Christopoulos, A., 61 Christova, Y., 145–146, 248, 257–258, 283f, 285–286, 287f, 289, 396–398, 408–409 Chu, P.H., 353–354, 359–360 Chug, H., 84–85 Church, G.M., 380–381 Church, W.R., 361–362 Cintra, G., 311–313, 319 Cipolat, S., 208–209 Citron, M., 160, 176 Clackson, T., 62 Claessens, J.G., 359–360 Clark, K.M., 128–129 Clark, R.J.H., 210 Clarke, D.M., 195–196 Clemens, M., 39 Clemmer, K.M., 239–240, 282–284, 400, 419–420 Climent, I., 311, 316 Cohen, A.S., 50–52 Cole, S.T., 311–313, 319 Collakova, E., 314 Collen, D., 197–198 Colombo, A., 130 Compernolle, G., 61–62 Condron, M.M., 130 Confavreux, C., 350–351 Cong, L., 371–372, 380–381 Connelly, S.M., 128–129 Connolly, M.D., 50–52

Author Index

Conradt, B., 135 Conrath, K.E., 61–62 Conzelmann, E., 352, 370, 374 Cooley, J.W., 208–225 Cooper, T.G., 355–356 Copeland, R.A., 209–210 Coquelle, N., 100–101, 116, 258–259, 268, 401, 409–410 Cordier, B., 100–123 Corringer, P.-J., 61 Corsmit, E., 196–197 Coughlan, S.J., 314 Court, D.L., 45 Cox, D., 380–381 Craessaerts, K., 128–129, 137, 186–187, 230, 296 Cranenburg, E.C., 386 Craney, A., 38–39 Cravatt, B.F., 112–114, 397 Creaser, S.P., 159–160 Crick, D.C., 311–314, 316, 319 Cromie, K.D., 86 Crosbie-Staunton, K., 85 Crown, D., 87 Crump, C.J., 159, 175 Cui, Y., 87 Cunniff, M.M., 371–372 Cupers, P., 128–129, 137, 186–187, 230, 296 Curnow, P., 264–265 Czech, C., 159, 163 Czirr, E., 159–160 Czogalla, K.J., 362–363, 378

D D’Haeze, W., 314, 316, 337 D’Huyvetter, M., 87–88 D’Mello, F., 70 Daelemans, D., 84–86 Dahan, S., 78 Dalbey, R.E., 36–54, 280–282 Dallner, G., 311, 316 Dang, S., 132–133, 136–138, 146f, 147 Danks, A.M., 159–160 Danziger, J., 386 Das, C., 198, 289–290 Das, P., 159–160 Date, T., 36

443 Datsenko, K.A., 320 Daugherty, P.S., 419, 421 Dave, U.P., 128–130, 296 Davidovic, L., 230–231, 279–280 Davidson, E., 84 Davie, E.W., 366 Davis, L., 27 de Boer-van den Berg, M.A., 356–357 De Bruyne, E., 87–88 De Genst, E., 61–62 de Gier, J.W., 256–258 De Haard, H.J., 86–88 De Jaegere, P., 87 de Jong, A., 36 de Jong, J.M., 39–40, 43 de Kruijff, B., 39–40, 43 De Luca, E., 313–314 De Metz, M., 355–356 De Pauw, P., 87–88 De Rosa, M., 314 De Strooper, B., 60–88, 128–129, 137–138, 143–145, 158–159, 186–187, 189–191, 193–194, 197–198, 208–209, 230–231, 296, 313–314 De Taeye, S., 81, 87 De Tavernier, E., 81, 87 De Winter, A., 61–62 De Wit, R.H., 86 Debnath, J., 311–313 DeBose-Boyd, R.A., 129–130 Declerck, P.J., 61–62 Decottignies-Le Marechal, P., 356–357 Delaney, R., 355–356 Delarue, M., 61 Dell, A., 355 DellaPenna, D., 314 Deluz, C., 61–62 Demel, R.A., 39–40, 43 Deng, Q.W., 87–88 Denisov, I.G., 65 Denninger, E., 313–314 Depla, E., 86–87 Desbois, C., 350–351 Descamps, F.J., 65–66 Desmyter, A., 61–62 Desvaux, C., 85 Detalle, L., 81, 86–87 Dev, I.K., 43

444 Devoogdt, N., 63–64, 75, 81, 87–88 Dewilde, M., 60–88 Dhiman, R.K., 311–313 DiCarlo, J.E., 380–381 Dickey, S.W., 8, 100, 135–136, 145–146, 230–231, 233–236, 239–240, 244–246, 248–250, 282–285, 287f, 288–290, 301–305, 396–398, 408 Diehl, T.S., 128–129, 137, 186–188, 198, 230, 289–290, 296 Digilio, G., 313–314 DiMauro, S., 311 Dimitrov, M., 144–145 DiMuzio-Mower, J., 128–129, 186–187, 194, 296–297, 399–400 Dmitriev, O.Y., 61 Dodd, R.B., 61, 128–129, 193–194 Dolios, G., 136–137, 158–159, 188, 298 Dombrecht, B., 81, 87 Dominguez, E., 112 Dominik, P.K., 65 Dong, J.X., 87 Doranz, B.J., 84 Dorner, T., 379 Dorner-Ciossek, C., 159–160 Doshi, R., 63–64 Dotsch, V., 138 Doublie, S., 330–331 Dougan, D.R., 340 Dougherty, W.G., 272–274 Dowhan, W., 414–415 Doyen, C., 80–81 Doyle, E.L., 372 Draheim, R., 102–104, 256–257 Dreier, J., 39–40 Dreier, T., 86–87 Drew, D., 256–258, 328–329, 333–334 Drittij-Reijnders, M.J., 353–354 Du, L., 83 Dube, M.-P., 313–314 D€ ubel, S., 85–86 Dubord, P., 313–314 Ducy, P., 350–351 Dumont, J., 361 Dumont, M.E., 128–129 Dumpelfeld, B., 171 Duncan, E.A., 129–130, 230

Author Index

Dunstan, C., 350–351 Durst, F., 101–102 Dutta, K., 214–216 Dutton, R.J., 368–369

E Eads, J.C., 209–210 Eagleburger, M.K., 210 Ebenezer, N., 313–314 Ebinu, J.O., 128 Ebke, A., 159, 171, 175–176, 240, 303–304 Eckman, J., 81 Edbauer, D., 137–138, 176, 187–188, 296–297 Edirisinghe, J.N., 313–314 Edwards, P.A., 316 Egan, W., 350–351, 355 Eimer, S., 167 € Ekici, O.D., 36–54, 280–282 Elad, N., 61, 68–69, 76, 85, 193–194 Elliott, G.R., 353–354 Ellman, J., 285–286, 287f Emde, A., 80–81 Emerson, S.U., 83–84 Engelbrecht, J., 46–47 Engelhardt, O.G., 87 Engelke, J.A., 356–357 Engelman, D.M., 208–209 Engen, J.R., 357–358 Engstr€ om, S., 336 Enos, M.D., 61 Erez, E., 128, 409 Eriksen, J.L., 159–160 Eriksson, N., 370 Ernster, L., 358–359 Esler, W.P., 186–187 Esmon, C.T., 355–357 Esnouf, M.P., 356–357 Espenshade, P.J., 129–130 Esposito, G., 313–314 Esselmann, H., 144–145 Estep, P., 81 Estes, S., 361 Esvelt, K.M., 380–381 Ettenberg, S.A., 87

445

Author Index

Euwart, D., 361 Evans, S., 313–314 Even-Desrumeaux, K., 75 Everaert, H., 87–88 Eylenstein, R., 83

F Fa´bry, M., 83 Fairlie, D.P., 282–284 Falcone, B., 397–398 Fan, Q.R., 101–102 Fan, Z., 60–61 Farner, K.C., 60–61 Fasco, M.J., 352–354, 358–360 Fass, D., 128, 131, 135–136, 145–146, 208–209, 231, 257–258 Federico, A., 313–314 Fedoriw, N., 128–129 Feng, G., 371–372 Feng, L., 2, 100, 130–131, 135–136, 145–146, 146f, 208–209, 231, 280–282, 297, 397–400 Feng, X., 311–313, 319 Fernandez, I.S., 148 Ferna´ndez, L.A´., 63–64 Fernandez, M.A., 60–61, 143–145, 298, 301–302 Fernlund, P., 350–351, 355 Ferraro, D.A., 80–81 Ferraro, J.R., 224 Ferron, M., 350–351 Feuerstein, G.Z., 361–362 Fields, G.B., 420–421 Fikes, J.D., 46–47 Filutowicz, M., 272–274 Findeis, M.A., 159–160 Finn, J., 340 Fischer, E.C., 63–64 Fischer, S.J., 102–104, 131, 135–136, 145–146, 231, 256–259, 280–282, 396 Fish, B.A., 175 Fleetwood, F., 63–64 Fleig, L., 129 Fluhrer, R., 130, 240, 296–297, 303–304 Flynn, J.M., 25 Foletti, D., 81 Fontes, F.L., 311–313, 319 Forsgren, M., 311, 316

F€ orster, T., 408–409 Fourel, I., 354, 370, 376–378 Fowler, B.J., 45 Fraering, P.C.., 138, 144–145, 159–160, 296–298 Fraile, S., 63–64 Francis, R., 187–188 Franzblau, S.G., 311–313 Fraser, P.E., 137–138 Fredericks, W.J., 313–314 Free, D.L., 314 Freeman, M., 100–101, 107–110, 117–118, 121, 128–129, 145–146, 208–214, 230–231, 239–240, 248, 256–258, 279–280, 282–286, 283f, 287f, 288–290, 296, 396–401, 408–409, 419–421 Fregin, A., 352, 362–363, 370, 374, 378 Freitas-Junior, L., 311–313, 319 Fre`re, J.M., 61–62 Freret, T., 60–61, 298 Frey, J.G., 49–50 Frickey, T., 316 Fridy, P.C., 78 Friedman, P.A., 355–356 Friedmann, E., 130 Frillingos, S., 189–191 Frisoni, G.B., 186–187 Fritsch, E.D., 105 Fr€ olich, N., 101–102 Frosch, M.P., 60–61 Fujisaki, S., 311 Fukuda, R., 65–66, 138 Fukumori, A., 120–121, 130, 158–181 Fukushima, N., 311, 337 Fukuyama, T., 194 Funamoto, S., 137, 143–144, 158–159 Furie, B.C., 356–357, 378–379 F€ urstenberg, A., 61 Fuwa, H., 194

G Gaffney, K., 100–101 Gainkam, L.O.T., 87–88 Galan-Diez, M., 350–351 Gale, N., 100, 257–259, 268, 396–397, 400–401, 419–420 Galleni, M., 61–62 Galley, G., 159–160

446 Galmozzi, A., 112 Gambacorta, A., 314 Ganglberger, E., 84 Gansler, J., 362–363, 378 Ganzke, T.S., 314 Garcia, K.C., 61 Garcia, M., 100–101, 107–110, 117–118, 230–231, 248, 256–258, 396, 398, 400 Gardell, S.J., 128–129, 399–400 Gardill, S.L., 352–353 Gardner, E.J., 355–356 Gartner, F.H., 402 Garvey, C.F., 129 Gaudry, M., 356–357 Gebhardt, O., 360–361 Gee, S.J., 87 Geiger, B., 129 Geisen, C., 379 Genet, C., 198, 289–290 Geng, Y., 101–102 Gennis, R.B., 311–313, 319 Gentry, P.R., 61 George, S., 313–314 Georges, G., 80–81 Gertsik, N., 175 Geweke, L.O., 356–357 Ghahroudi, M.A., 61–62 Ghasriani, H., 100–101, 112–114 Gheesling, L.L., 366 Giagounidis, A., 80–81 Gibson, F., 311, 316 Gilbert, B.E., 86–87 Gildehaus, F.J., 171 Gimpl, K., 257–258, 265, 427 Ginalski, K., 230–231 Ginsburg, D., 379 Girardot, J.M., 355–356 Goel, S., 87–88 Goetz, R.M., 258–259 Golde, T.E., 130, 159 Goldstein, J.L., 2, 36, 128–130, 230, 296 Gong, X., 132–133, 136–138, 146f, 147, 358–359 Gonzales, V., 186–187 Gonzalez-Pajuelo, M., 87 Goodall, J.J., 39–40 Goodman, H.L., 356 Goodman, J.M., 36

Author Index

G€ orlich, D., 84–85 Goto, N.K., 100–101, 112–114 Gottstein, D., 138 Gouaux, E., 329 Goud, B., 66–67 Grammer, G., 130 Grasso, L., 61–62 Green, M.R., 16 Griffin, P.R., 39–40 Griffiths, A.D., 70 Grigorenko, A.P., 130 Grim, M.G., 176 Grinkova, Y.V., 65 Grinnell, B.W., 361 Grishin, N.V., 230–231 Groenen-van Dooren, M.M., 352–353 Gross, C.A., 2–4 Growney, J.D., 87 Gruenbaum, Y., 135 Grunberg, J., 163, 176 Grunberger, K., 84 Gr€ unler, J., 311, 316 Gu, Y., 8, 240, 296–298, 301, 303, 305 Guardiola-Serrano, F., 130 Gubbens, S., 148 Guell, M., 380–381 Guernsey, D., 313–314 Guhde, G., 137 Gulati, A., 311–313, 319 G€ unnewich, N., 337 Guo, M.S., 2–4 Guo, R., 100–101 Guo, R.T., 311–313, 319 Gutala, R., 358–359 Gutsmiedl, A., 159–160

H Ha, Y., 2, 100–101, 112–114, 116, 131, 135–136, 145–146, 208–209, 231, 257–258, 265, 280–282, 285–286, 287f, 297, 396–397, 409 Haagen, Y., 340–341 Haapasalo, A., 158 Haass, C., 130, 137–138, 143, 158–160, 162, 167, 171, 175–176, 187–188, 296–297 Habib, N., 380–381 Hacker, D.L., 144–145 Hackmann, Y., 101–102

Author Index

Haddad, D.M., 313–314 Hadravova´, R., 100–101, 112–114, 285–286 Haffner, C., 130 Hageman, J.M., 356–357 Hagemans, D., 283f Hagen, W., 61, 76, 85, 193–194 Hagihara, Y., 78 Hale, J.E., 356–357 Hall, S.E., 314 Hallgren, K.W., 359–361 Halsey, C.M., 210 Hamakubo, T., 65–66 Hamamoto, M., 311 Hamann, T., 65 Hamilton, J.A., 311, 316 Hammarstrom, L., 86–87 Hammed, A., 354, 370, 376–378 Hamze, M., 256–257 Handford, J.I., 45 Hanes, J., 63–64 Haque, J.A., 362–363, 369 Harada, I., 210 Hardy, J., 158–159, 186–188 Hardy, J.A., 137 Harmsen, M.M., 86–88 Harrington, D.J., 377–378 Harris, B.Z., 129–130 Harris, P.W., 36, 38 Harrison, B.A., 159 Hartenian, E., 382 H€artle, S., 83 Hasan, M.T., 2, 128, 230, 296 Hasegawa, M., 320, 401, 421 Hashimoto, T., 60–61 Hass, M., 359–360 Hassaine, G., 61–62 Hassanzadeh-Ghassabeh, G., 61–62, 87–88 Hatanaka, Y., 194 Hauben, E., 130 Haug-Kroper, M., 130 Haustraete, J., 81 Hay, F.C., 83 Hayashi, I., 65–66, 137–138, 187–188, 296–297 He, M., 63–64 He, W., 136–137 Healey, J.F., 80–81 Hebling, C.M., 357–358

447 Hederstedt, L., 314–315 Hedstrom, L., 280, 289 Heemskerk, J., 87–88 Hegarty, J., 313–314 Heiberg, A.C., 370 Heide, L., 310–311, 316, 319, 323–324, 340–341 Heinemann, U., 138 Helfrich, M., 315–316 Hemker, H.C., 355–356 Hemmi, H., 314 Henderson, R., 282 Hendrickx, M.L.V., 61–62 Hendrix, H., 356–357 Hengartner, M.O., 136 Henn, W., 313–314 Herl, L.D., 60–61 Hermanson, G.T., 402 Herms, J., 171 Herreman, A., 197–198 Hersh, B.M., 135 Hersh, E.M., 49 Hester, R.E., 210 Heukers, R., 87–88 Higel, F., 130 Higgins, B., 313–314 Higgins, G.A., 137 Higgins-Gruber, S.L., 378–379 Higo, T., 159, 175, 189–191, 194–195 Hikita, C., 14 Hildebrandt, E.F., 360–361 Hillerich, B.S., 340 Hino, T., 65 Hinshaw, J.C., 311–313 Hirano, M., 311 Hiromoto, K., 194 Hirota, Y., 313–314, 316 Hirotani, N., 158–159, 188 Hitz, W.D., 314 Hizukuri, Y., 2–32 Hjelm, A., 256–258 Hjelmeland, L.M., 265–266 Hmila, I., 81 Ho, Y.K., 129–130 Hobson, S., 171 Hodroge, A., 354, 370, 376–378 Hoffmann, R., 419 Hofmann, K., 189–191

448 Holland, S., 313–314 Holm, L., 340 Holmes, O., 298–300 Holmgren, A., 352–353 Holsters, M., 314, 316, 337 Holz, J.B., 87 H€ olzer, W., 61–62 Hoogenboom, H.R., 70 Hope, G., 130 Horanyi, P.S., 128–129 Hori, H., 337 Horikoshi, Y., 158–159, 188 Horne, R.W., 238–239 Horre, K., 189–191 Hortnagel, K., 352, 370, 374 Horvitz, H., 135 Horvitz, H.R., 136 Hosfield, D.J., 340 Hou, Y.N., 87–88 Hovius, R., 61–62 Howard, C.R., 70 Howard, E.D., 272–274 Howard, J.B., 350–351 Hruby, V.J., 49 Hu, L., 256–257, 397–398 Hu, N.J., 333–334 Hua, X., 129–130, 230 Huang, C.Y., 223 Huang, H., 314–316, 319, 330, 334, 337 Huang, L., 87–88 Huang, M., 136–137 Huang, N., 63–64 Huang, Q., 128–129, 186–187, 194, 296–297, 399–400 Huang, T.Y., 353–354, 359–360 Huang, X., 100–101 Huang, Y., 258–260 Huang, Z.J., 408 Huault, S., 130 Hubbard, S.J., 208–209 Hudspeth, M.E., 311–313, 316 Huet, H.A., 87 Hufton, S.E., 87 Hultberg, A., 86–87 Hunerberg, M., 370 Hung, A.Y., 160 Hung, J.H., 285–286 Hunte, C., 266–267, 414–415

Author Index

Hutson, S.M., 352–353, 360–361 Hutton, M., 130 Huyck, L., 86–87 Hyman, B.T., 60–61

I Ibraheem, A., 409 Ihara, Y., 137, 143–144, 186–187 Ikeda, K., 379 Ikeda, M., 128–129, 186–187 Ikeya, T., 138 Ilagan, M.X., 137, 158 Imamura, Y., 194 Impens, N., 87–88 Imtong, C., 84 Inaba, K., 2–4 Inada, T., 45 Ingalls, J., 84 Ingram, B.O., 358–359 Inoue, N., 78 Inoue, S., 379 Inouye, M., 41, 43 Iqbal, U., 85 Ishihara, S., 143–144, 158–159 Isoo, N., 65–66 Israel, L., 130 Ito, K., 2–8, 10, 18–20, 25, 29–32, 45, 120–121, 210–212, 230–231, 256–257, 396–400 Ito, Y., 78 Ivanov, A., 60–61 Ivaskevicius, V., 352, 370, 374 Iwata, S., 265–266, 328–329, 333–334 Iwatsubo, T., 65–66, 137–138, 143, 158, 186–187, 189–198, 192f, 195f

J Jacobs, M.R., 356–357 Jacobsen, H., 159–160, 163, 171 Jacobsen, H.J., 80–81 Jacobson, M.P., 340 Jahnen-Dechent, W., 357 J€ahnichen, S., 87 Jaiswal, A.K., 358–359 James, M.N., 36, 40, 131, 135–136, 145–146, 256–259, 280–282, 396 James, M.N.G., 102–104, 231, 280–282 Jamnani, F.R., 80–81

449

Author Index

Jansen, K., 130 Janssen, C.A., 353–354 Janssen, Y.P., 352–353 Jarrett, J.T., 186–187 Javitch, J.A., 189–191 Jaworska, A., 171 Jeffrey, P.D., 2, 131, 136, 146, 146f, 297 Jegersch€ old, C., 86 Jenkins, N.A., 186–187 Jensen-Jarolim, E., 84 Jiang, H., 313–314 Jiang, S., 83 Jiang, S.J., 285–286 Jiang, T., 311–313, 319 Jidenko, M., 101–102 JiJi, R.D., 208–225 Jin, D.Y., 352, 357–361, 364–365, 367–370, 372, 374, 377–378, 381–386 Jin, L., 83 Joedicke, L., 101–102 Johan, L., 352–354, 360–361 Johansen, K., 86–87 Johansson, L.C., 336 Johnson, B.C., 355–356 Johnson, D.S., 159 Johnson, J.A., 87 Johnson, K., 186–187 Johnson, R.B., 36 Johnston, S.A., 272–274 Jones, J.P., 355–356 Jones, S.E., 70 Jordan, T., 209–210 Jorgensen, M.J., 356–357 Jorgenson, J.W., 357–358, 378–379 Jost, M., 340–341 Ju, H., 84 Jucker, M., 159–160 Julius, D., 148 Jumpertz, T., 175 Jung, J., 314 Jung, J.I., 159 Junge, F., 101–102 Jutte, H., 46–47

K Kaback, H.R., 189–191 Kachala, M., 83 Kaesler, N., 357

Kaether, C., 137, 167 Kahn, P.C., 41, 43 Kahne, D., 397–398 Kainou, T., 311 Kajava, A.V., 46–47 Kakkar, E., 84 Kaltak, L., 129 Kamiya, Y., 311 Kamp, F., 240, 303–304 Kampani, K., 84 Kan, T., 194 Kanehara, K., 2–8, 10, 25, 230–231 Kang, H.J., 256–258 Kania, M., 39–40 Kappel, W.K., 357 Kapust, R.B., 272–274 Karamyshev, A.L., 46–47 Karamysheva, Z.N., 46–47 Karla, A., 37–41, 47 Karlin, A., 189–193 Karpowich, N.K., 258–259 Karran, E., 186–187 Karsenty, G., 350–351 Karunanandaa, B., 314 Kasahara, M., 328–329, 333–334 Katona, G., 336 Katzenmeier, G., 84 Kawai, M., 356 Kawamukai, M., 311 Kawate, T., 329 Kazmierski, W.M., 49 Ke, N., 310–341 Keegan, S., 78 Keller, S., 257–258, 265, 340–341, 427 Kellogg, B.A., 340 Kemmerling, M., 314 Keshavarz-Moore, E., 71 Keyaerts, M., 87–88 Khalimonchuk, O., 316 Khan, K., 285–286 Khvorova, A., 352, 370 Kidd, D., 112 Killian, J.A., 39–40 Kim, G., 195–196 Kim, H., 333–334 Kim, H.J., 316 Kim, M., 100–101, 314 Kim, M.J., 259–260

450 Kim, Y.S., 208–209 Kimberly, W.T., 128–129, 186–188, 230, 296–297 Kinch, L.N., 230–231 King, C.C., 45 Kinne, J., 61–62 Kinney, A.J., 314 Kintner, C., 128 Kirfel, G., 359–360 Kirkin, V., 130 Kisiel, W., 366 Kiso, Y., 189–191 Kitamura, T., 197–198 Kiyose, N., 78 Kjær, I., 80–81 Klafki, H.W., 144–145 Klapper, D.G., 46–47 Klco, J.M., 195–196 Klein, C., 80–81 Klein, T.E., 370 Kleinridders, A., 358–359 Kleinschmidt, J.H., 65 Klenk, D.C., 402 Klenotic, P.A., 41, 45 Klepsch, M.M., 102–104, 256–257 Klutkowski, J.A., 60–61, 298 Knapen, M.H., 386 Knapp, R.J., 49 Knauer, T.E., 353 Knight, C.G., 420–421 Knight, W.B., 39–40 Kn€ obl, P., 87 Knobloch, J.E., 356–357 Kobilka, B.K., 61 Kodama, T., 65–66 Kodama, T.S., 158–159 Koefoed, K., 80–81 Koenders, M., 87 Koenig, J.L., 210 Koga, Y., 314 K€ ogel, J., 315 Koide, A., 63–64 Koide, K., 18–20, 29–32, 120–121, 208–209 Koide, S., 63–64 Koide, T., 361–362 Kok, R.J., 87–88 Kokado, Y., 311

Author Index

Komuro, R., 128–130, 296 Koo, E.H., 60–61, 143, 159–160 Koonin, E.V., 230–231, 279–280 Kopan, R., 36, 128–129, 137–138, 158, 208–209 Kornhuber, J., 144–145 Koshino, Y., 197–198 Kossiakoff, A.A., 65 Kostka, M., 187–188 Koszelak-Rosenblum, M., 128–129 Koth, C.M., 100–101, 107–110, 117–118, 230–231, 248, 256–258, 396, 398, 400 Kounnas, M.Z., 159–160 Kovacs, D.M., 158 Koyama, A., 197 Kra´l, V., 83 Krause, M., 313–314 Kremer Hovinga, J.A., 87 Kremmer, E., 130, 162, 167, 176 Kress, J., 49–50 Kretner, B., 158–181 Kreuzman, A.J., 39 Krieg, P.A., 16 Krishnaswamy, S., 381–382 Kristensen, J.V., 100–101, 231 Krohn, R.I., 402 Kroos, L., 2–4 Kruger, T., 357 Kruk, J., 310 Kruse, A.C., 61 Kruth, H.S., 313–314 Krzysiak, A.J., 143–145, 301–302 Ksenzenko, V.N., 46–47 Kshirsagar, R., 361 Kubicek, J., 107–110 Kufka, J., 337 Kuhn, A., 46–47 Kuhn, P.H., 130, 176 Kuivaniemi, H., 313–314 Kukar, T., 159 Kulanthaivel, P., 39 Kulkarni, K., 128–129 Kulman, J.D., 362–363, 369 Kumar-Singh, S., 196–197 Kume, H., 176, 186–187, 197 Kummel, D., 138 Kunji, E., 258 Kunzel, U., 130

Author Index

Kuo, D.W., 39–40 Kuperman, A.A., 379 Kurosu, M., 311–313 Kutsunai, S.Y., 316 Kuzuyama, T., 340–341

L Labahn, J., 107–110 Labelle, E., 313–314 Lada, B., 208–225 Ladd, T.B., 159 Ladlow, M., 49–50 Laeremans, T., 64–67, 70–72, 80 Lahoutte, T., 85, 87–88 Lai, C.H., 285–286 Lai, M.T., 128–129, 186–187, 194, 296–297, 399–400 Lake, S., 311, 316 Lal, P., 313–314 Lam, K.S., 49 Lammens, A., 80–81 Lammich, S., 137 Lamoureux, J.S., 145–146, 258–259 Langosch, D., 8, 128, 285 Lansbury, P.T., 186–187 Lapecorella, M., 350–351 Large, J., 286, 287f, 289, 396–398, 408–409 Larson, A.E., 355–356 Lassere, T.B., 195–196 Lassner, M.W., 314 Lattard, V., 354, 370, 376–378 Laue, K., 350–351 Lauwereys, M., 61–62, 86–87 LaVoie, M.J., 187–188, 296–298 Law, C.J., 258–259 Lawson, D.M., 340–341 Lazareno-Saez, C., 100–101, 116, 145–146, 258–259, 268, 401, 409–410 Lazarov, V.K., 138 le Maire, M., 101–102 Lechartier, B., 311–313, 319 Lednev, I.K., 210, 223 Lee, C., 256–258, 333–334 Lee, C.-T., 84–85 Lee, C.-W., 63–64 Lee, J.I., 45–47 Lee, J.J., 352

451 Lee, J.R., 121, 129, 208–209, 211–212, 230, 248, 279–280, 285–286, 296, 398, 400 Lee, K., 314 Lee, M.K., 186–187, 195–196 Lee, M.T., 370 Lee, S.M., 314 Leeds, J.A., 2–4 Lefkowitz, R.J., 85–86 Lehrman, S.R., 356–357 Lei, X., 258 Leighton, B.H., 189–191 Lemaire, M., 87–88 Lemberg, M.K., 8, 100–123, 128–130, 187–188, 211–212, 230–231, 248, 256–258, 285, 297, 396–398, 400 Lemieux, J., 396–433 Lemieux, M.J., 100–104, 116, 131, 135–136, 145–146, 208–209, 231, 256–274, 280–282, 396–397, 400–401, 409–410, 414–415, 419–420, 427 Lenaerts, A.J., 311–313 Lentz, B.R., 357–358 Lenz, S., 315 Leppik, R.A., 311, 316 Leppla, S.H., 87 Lepsˇ´ık, M., 100–101, 145–146, 281f, 282–285, 283f, 287f, 288–290, 396 Lerch, M., 258 Letellier, M.C., 230–231, 279–280 Leuchtenberger, S., 159–160 Leung, J., 333–334 Leuther, K.K., 272–274 Levchenko, I., 25 Levesque, G., 128–129, 186–187 Levin, E.J., 315–316, 319, 330, 334, 337 Levitan, D., 187–188 Levy, D., 235–236 Levy-Lahad, E., 208–209 Leysath, C.E., 87 Li, D., 138 Li, H., 136–138 Li, J., 83, 87 Li, K., 311–313, 319 Li, S.-M., 311, 337, 340–341 Li, T., 79–80, 87–88, 208–209, 352, 370 Li, W., 310–341, 352–353, 368–369 Li, W.P., 129–130

452 Li, X., 2, 60–61, 100, 132–133, 136–138, 146f, 147–148, 158, 208–209 Li, X.D., 259–260 Li, Y., 61, 78, 84, 193–194, 311–313, 319 Li, Y.-M., 128–129, 159, 186–187, 194, 258, 296–297, 399–400 Liang, Y., 128–129, 186–187 Liao, J., 420–421, 431–434 Liao, M., 148 Liapakis, G., 189–191 Libert, C., 81 Lichtenthaler, S.F., 130, 158–159 Lill, C.M., 60–61 Lillelund, O.K., 100–101, 231 Lim, W.A., 129–130 Lima, S., 2–4 Lin, P.J., 352, 370, 378–379 Lin, S., 380–381 Lind, C., 358–359 Lindorff-Larsen, K., 100 Lindsley, C.W., 61 Lisch, W., 313–314 Liskamp, R.M.J., 100–101, 285–289, 287f, 396 Lismont, S., 60–61, 68–69, 76, 85, 159–160, 193–194 Liu, J., 39–40, 87–88 Liu, S., 310–341 Liu, S.K., 285–286 Liu, W., 335–337 Liu, X., 285–286, 287f Liu, Y., 81, 112–114, 420–421, 431–434 Liu, Y.L., 311–313, 319 Liu, Z., 84 Lively, M.O., 37–41, 47 Lleo, A., 60–61, 159–160 Lockless, S.W., 315–316, 319, 330, 334, 337 Lodhia, P., 285–286 Loetscher, H., 163 L€ ofblom, J., 63–64 Lohi, O., 398 Lohr, F., 138 Lollar, P., 80–81 Lombard, C., 420–421 Loo, T.W., 195–196 Lo´pez, L.C., 311 Lord, R.C., 210 Lorenz, H., 128–129

Author Index

Lorsch, J., 107–110 L€ ow, C., 86 Lowman, H.B., 62 Lu, C.Z., 2–4 Lu, P., 2, 101–102, 142f, 144–145, 146f, 147, 189, 191–193, 297 Lu, S.H., 193–194 Lu, S.H.J., 61 Luckerath, K., 130 Luebbers, T., 159–160, 171, 175–176 Luke, I., 45 Lundstrom, J., 352–353 Luo, C., 39–40 Luo, Y., 85 Lupas, A.N., 310, 316–319, 340–341 Lutsenko, S., 61 Lynas, J.F., 289

M Ma, D., 2, 101–102, 142f, 144–145, 146f, 147, 189, 191–193, 297 Mack, D.O., 355–356 Madahar, V., 420–421, 431–434 Madala, P.K., 208–209, 282–284 Maegawa, K.-I., 2–4 Maegawa, S., 18–20, 120–121, 210–212, 256–257, 282–284, 396–400 Maertens, B., 107–110 Maes, E., 79–80 Magdeleyns, E.J., 386 Magnusson, S., 355 Mahapatra, S., 311–313 Mahboudi, F., 80–81 Mahfoud, O.K., 85 Mahurkar, S., 313–314 Major, D., 87 Mak, M.W., 100, 145–146, 257–259, 268, 396–397, 400–401, 419–420 Makarova, K.S., 230–231, 279–280 Mali, P., 380–381 Mali, W., 87–88 Malik, A.A., 84 Malkowicz, S.B., 313–314 Mallia, A.K., 402 Mancia, F., 136–137 Maneewatch, S., 84 Manglik, A., 61 Maniatis, T., 16, 105

Author Index

Manolaridis, I., 128–129, 208–209 Manzari, M.T., 80–81 Marcadier, J., 313–314 Marcotte, H., 86–87 Margolles, Y., 63–64 Mariani, G., 350–351 Marı´n, E., 63–64 Marinova, M., 359–360 Markovitz, R.C., 80–81 Marks, J.D., 70 Marschall, A.L.J., 85–86 Martin, L., 130 Martin, L.F., 353–354, 358–361 Martin, M.M., 143–145, 301–302 Martı´nez, V., 63–64 Martı´nez-Arteaga, R., 63–64 Martinhago, C.D., 383–386 Martoglio, B., 2, 117–118, 128–130, 187–188, 230, 297 Maruyama, K., 176, 186–187, 197 Mas, V., 86–87 Matagne, A., 61–62 Matagrin, B., 354, 370, 376–378 Matasci, M., 144–145 Mathews, T.P., 61 Matschiner, J.T., 352–353 Matsuda, H., 311 Matsuo, E., 2–4, 7, 11, 14 Matsuyama, S., 14, 17 Matthys, P., 85 Matuschek, M., 340–341 Maussang, D., 65–66, 87 May, P.C., 84 Mayer, C., 316–319 Maylandt, K., 130 McAuliffe, J.C., 70 McCafferty, J., 70 McClure, D.B., 361 McConlogue, L., 160 McDonald, M.G., 362–363, 369 McGarvey, T., 313–314 McGrath, G., 187–188 McKee, T.D., 159–160 McLauchlan, J., 130 McMullan, G., 148 McNeil, M.R., 314, 316, 337 Medana, C., 313–314 Meeks, S.L., 80–81

453 Meganathan, R., 311–314, 316 Mei, B., 361 Meijer, P.J., 80–81 Meissner, C., 128–129 Meldal, M., 50–51 Mello, C.C., 130 Melton, D.A., 16 Melzer, M., 311, 316, 319, 323–324 Menendez, J., 100–101, 107–110, 117–118, 230–231, 248, 256–258, 396, 398, 400 Menu, E., 87–88 Mera, P., 350–351 Metzger, U., 340–341 Meyn, L., 176 Mielke, J., 176 Mikelsaar, R.-H., 86–87 Mikhonin, A.V., 209–210, 209f, 221–222 Mikkelsen, T.S., 382 Miles, A., 81 Miller, J.A., 355–356 Minehira, M., 311 Mio, K., 138 Miroux, B., 102–104, 256–257, 326–328 Misik, A., 100–101, 107–110, 117–118, 230–231, 248, 256–258, 396, 398, 400 Mitchell, D.A., 311–313, 319 Mito, K., 337 Mitsky, T.A., 314 Miyashita, H., 138 Miyazaki, N., 78 Mizuno, S., 2–7, 27–32 Mizusawa, H., 186–187 Mizushima, S., 14, 17 Mlynarczyk, K., 138 Moayeri, M., 87 Moerner, W.E., 61 Mogi, T., 337 Mohamed, B.M., 85 Mohr, K., 412–414, 428–430 Moin, S.M., 8, 100, 211–212, 235, 282, 285, 289, 299–300, 397–398 Moliaka, Y.K., 130 Møller, J.V., 101–102 Molohon, K., 311–313, 319 Mondou, M.-H., 78 Mongrain, V., 313–314 Montagna, D.R., 8, 240, 296–298, 301, 303, 305

454 Moomaw, J., 36 Moon, J.-K., 314 Mooney, P., 148 Moore, B.D., 159 Moore, C.L., 186–187 Morais, V.A., 313–314 Moran Luengo, T., 283f Moreau, C., 354, 370, 376–378 Morgan, C.R., 357–358 Mori, H., 2–7, 11, 14, 27–32, 320, 401, 421 Mori, K., 158–159 Morii, H., 314 Morishima, K., 189–191, 193–194 Morishima-Kawashima, M., 143–144, 158–159, 188 Moriyama, Y., 328–329, 333–334 Morizzo, E., 81, 87 Mormino, E., 186–187 Morohashi, Y., 189–198 Morris, D.P., 378–379 Morris, H.R., 355 M€ ossner, E., 80–81 Mosyak, L., 101–102 Moutel, S., 66–67 Mugoni, V., 313–314 Mujic-Delic, A., 65–66, 86 Muller, C.R., 370 Muller, J., 359–360 Muller, K., 419 Mumford, A.D., 377–378 Mumm, J.S., 128–129, 137, 186–187, 230, 296 Munter, L.M., 256–257 Muroya, A., 311, 337 Murphy, M.P., 313–314 Murray, C.J., 70 Murray, M., 84 Murrey, H.E., 175 Mutucumarana, V.P., 357, 378–379 Muyldermans, S., 61–66, 70–71, 81, 83–85, 87–88

N Nagashima, Y., 143–144, 158–159 Naik, R.R., 70 Naini, A., 311 Nakada-Nakura, Y., 65 Nakagawa, K., 313–314, 316, 379

Author Index

Nakagawa, T., 311 Nakajima, H., 197–198 Nakamoto, K., 224 Nakamura, Y., 45 Nakayama, T., 314 Nall, T., 282–284 Nam, Y., 330 Nandi, D.L., 352–353 Napolitano, M., 350–351 Narayanan, S., 137 Narayanasamy, P., 311–313 Narlawar, R., 159 Nasser-Ghodsi, N., 60–61 Natsugari, H., 194 Nauman, C., 81 Nelsestuen, G.L., 350–351 Nesmeyanova, M.A., 46–47 Ness, J., 175 Neuhaus, G., 314, 316 Neuhoff, V., 120 Neutze, R., 336 Newman, P., 350, 386 Newstead, S., 328–329, 333–334 Nguyen, A.W., 419, 421 Ni, C.Y., 208–209 Nichols, A.J., 361–362 Nickerson, M.L., 313–314 Niederfellner, G., 80–81 Nielsen, H., 46–47 Nielsen, R.C., 101–102 Niewold, T.A., 86–87 Niimura, M., 137–138, 187–188, 296–297 Nijhawan, D., 2, 128, 230, 296 Ning, Q., 84 Nishimura, M., 187–188 Nishimura, O., 2–4, 7, 11, 14 Nishimura, Y., 311 Nishino, T., 311, 314 Nissen, P., 101–102 Nizak, C., 66–67 Nji, E., 256–258, 333–334 No, J.H., 311–313, 319 Noel, J.P., 340–341 Nogi, T., 2–7, 27–32 Nohturfft, A., 129–130 Nojiri, H., 311 Nomura, N., 65 Nomura, Y., 65

455

Author Index

Nordlund, P., 86 Nørholm, M.H.H., 63–64 Nosaka, T., 197–198 Novak, P., 43 Nowicka, B., 310 Noy, P.J., 285–286 Nozaki, K., 311 Nudelman, I., 78 Nukina, N., 186–187 Nury, H., 61–62 Nyborg, A., 130

O Oberstein, A., 130–131, 135–136, 145–146, 146f, 231, 280–282 Oda, T., 2–4, 22–25 Odaka, A., 186–187 Odam, E.M., 353–354 Oestereich, F., 256–257 Offord, R.E., 356–357 Ogasawara, S., 128–129 Ogura, T., 138 Ohara, K., 311, 337 Ohki, Y., 159, 175, 189–191, 194–195 Oi, R., 2–4, 22–25 Okada, K., 311 Okamura, Y., 194 Okano, T., 313–314, 316, 379 Oki, T., 197–198 Okochi, M., 137, 158–159, 167 Okuda, N., 313–314, 316 Okumura, Y., 320, 401, 421 Oladepo, S.A., 209–210 Oldenburg, J., 352, 358–360, 362–363, 370, 374, 378–379 Oldfield, E., 311–313, 319 Oliveira, S., 87–88 Olivo, J.C., 176 Olson, R.E., 355–356 Oltersdorf, T., 160 Omote, H., 328–329, 333–334 Omura, S., 361–362 Onufryk, C., 2–4 O’Reilly, R.A., 359 Orlean, P., 311–313, 319 Orr, A., 313–314 Osawa, S., 159, 175, 188–191, 193–196

Osenkowski, P., 138, 143–145, 189, 298, 301–302 Ostaszewski, B.L., 128–129, 186–188, 230, 296–298 Oster, U., 315 Ostrom, L., 87 Otto, M., 144–145 Otzen, D.E., 100 Overall, C.M., 284 Owen, J., 352, 360–361

P Pacanowski, M., 370 Paes, C., 84 Paetzel, M., 36–41, 43, 45, 47, 280–282 Page, M.G., 39–40 Page, R.M., 159–160, 171 Painter, P.C., 210 Palmer, T., 45 Palmier, M.O., 430–431 Palomo, C., 86–87 Pan, J.J.-J., 340 Pan, L.C., 356–357 Panavas, T., 216–217 Pan-Hammarstrom, Q., 86–87 Panigrahi, R., 256–274, 427 Panneels, V., 101–102 Pant, N., 86–87 Panwar, P., 100, 102, 257–259, 265, 268, 396–397, 400–401, 419–420, 427 Pardon, E., 63–64, 66–67, 71–72, 80, 85–86 Park, S.-R., 314 Parker, C.H., 357–358 Parker, E.T., 80–81 Parks, T.D., 272–274 Pascal, S.M., 214–216 Paschkowsky, S., 256–257 Paslawski, W., 100–101, 231 Patel, A., 361–362 Patricelli, M.P., 112–114 Patskovsky, Y., 340 Patterson, J.L., 360–361 Paturi, S., 298–300 Paul, S., 144–145 Peclinovska´, L., 100–101, 145–146, 281f, 282–285, 283f, 287f, 288–290, 396 Pedersen, M.W., 80–81 Peery, R.B., 36

456 Pei, D., 36–54 Pellegrini, L., 230–231, 279–280 Pellis, M., 63–64 Pelz, H.J., 352, 370, 374 Peng, S.B., 36, 39 Perez, F., 66–67 Perez-Revuelta, B.I., 159–160, 171 Pesold, B., 137–138, 176, 187–188, 296–297 Peters, J.A., 60–61 Petersen, J., 80–81 Petersen, T.E., 355 Pettersson, M., 175 Peyvandi, F., 87 Pezacki, J.P., 100–101, 112–114 Philipp, U., 176 Piedra, P.A., 86–87 Pierrat, O.A., 286, 287f, 289, 396–398, 408–409 Pietrzik, C.U., 159–160, 175 Pinero, G., 350–351 Pirici, D., 196–197 Pleiner, T., 84–85 Plikaytis, B.D., 366 Pl€ uckthun, A., 63–64 Poitevin, F., 61 Ponting, C.P., 130 Poole, S., 87 Popot, J.-L., 65 Post-Beittenmiller, D., 314 Postel, R., 313–314 Poulter, C.D., 314, 316, 340 Poungpair, O., 84 Powers, M.E., 45 Pozdnyakov, N., 175 Pradeep, C.-R., 80–81 Prentice, C.R., 350–351 Presnell, S.R., 357–358 Preusch, P.C., 352–353, 360–361 Prevost, M.S., 61 Price, A.R., 159 Price, P.A., 356–357 Prina-Mello, A., 85 Principe, L.M., 353–354, 358–360 Prokop, S., 137–138, 176 Provenzano, M.D., 402 Provost, S., 313–314 Prydz, H., 360–361

Author Index

Pryor, E.E., 128–129, 208–209 Prywes, R., 128–130, 296 Puime`ge, L., 81 Pujari, V., 311–313, 319 Purcell, R.H., 83–84 Purich, D.L., 405 Put, S., 85 Puthiyaveettil, R., 313–314

Q Qamar, S., 61, 193–194 Qi, Q., 314 Qi, S., 87–88 Qian, W., 360–361 Qi-Takahara, Y., 158–159, 188 Quintero-Monzon, O., 143–145, 301–302 Quinzii, C.M., 311 Quistgaard, E.M., 86 Qureshi, A.A., 256–258

R Radauer, C., 84 Rahbarizadeh, F., 80–81 Rajendra, E., 61, 189, 193–194 Rajpal, A., 81 Rakovich, T.Y., 85 Ramamoorthy, G., 340 Ran, F.A., 380–381 Rand, K.D., 357–358 Rao, G., 311–313, 319 Rapoport, T.A., 330, 352–353, 368–369 Rase, B., 358–359 Rasmussen, S.G.F., 64, 66–67, 71–72, 80, 85–86 Rather, P.N., 239–240, 282–284, 400, 419–420 Ratovitski, T., 186–187 Rausch, F., 337 Rawlings, N.D., 129–130 Rawson, R.B., 2, 36, 128–130, 208–209, 230, 296 Ray, P.H., 43 Raymond, R.M., 379 Razzano, P., 361 Rebagliati, M.R., 16 Reckel, S., 101–102 Regula, J.T., 137–138, 187–188, 296–297 Reid, F., 81

457

Author Index

Reiss, K., 130 Reith, M.E., 258–259 Reithmeier, R.A., 266–267, 414–415 Remmert, M., 316–319 Ren, F., 311–313, 319 Rennhack, A., 175 Rettie, A.E., 362–363, 369 Reutelingsperger, C.P., 350–351 ˇ eza´cˇova´, P., 83 R Rial, D.V., 269 Ribba, B., 80–81 Rich, D.H., 356–357 Richard, S.B., 340–341 Rieger, M., 81, 87 Riemer, D., 135 Rigaud, J.L., 235–236 Rikong-Aide, H., 356–357 Ring, A.M., 61 Ripp, K.G., 314 Rishavy, M.A., 359–361 Risley, P., 87 Ritchie, T.K., 65 Roach, C.A., 209–210 Robak, T., 80–81 Roberts, T.C., 39, 45 Robinson, R.A., 411–412, 426 Roby, P., 78 Rodgers, K.R., 209–210 Roepstorff, P., 350–351, 355 Rogaev, E.I., 128–130, 186–187, 208–209 Rogaeva, E.A., 128–129, 186–187 Rogozin, I.B., 230–231, 279–280 Rohrig, S., 163 Romesberg, F.E., 38–40, 45 Romig, H., 176, 187–188 Romiti, S., 357 Roobrouck, A., 81, 87 Roodt, J., 87 Roovers, R.C., 87–88 Rosalia, E.K., 80–81 Rosano, G.L., 272–274 Rose, K., 356–357 Rosen, H., 397 Rosenstr€ om, P., 340 Ross, C.M., 136–137 Rosseels, V., 87 Rossenu, S., 87 Rost, S., 352, 362–363, 370, 374, 378

Rottiers, P., 86–87 Rottnek, J.M., 314 Rotzer, C., 171 Rouleau, N., 78 Roussel, P., 39–40 Rout, M.P., 78 Routzahn, K.M., 272–274 Roy, A., 102, 107–110 Ruano-Gallego, D., 63–64 Ruddy, D.A., 187–188 Rudiger, S.G., 283f R€ udiger, W., 315–316 Ruf, A., 64, 66–67, 71–72, 80 Ruiz, N., 397–398 Runge, K.W., 359–361 Ryckaert, S., 63–64

S Sadowski, J.A., 353, 355–359 Saerens, D., 64–66, 70–71, 81 Saez, E., 112 Saffrich, R., 176 Saftig, P., 128–130, 137, 186–187, 230, 296 Sagi, S.A., 159–160 Sahasrabudhe, P., 129 Sahin-Toth, M., 189–191 Saido, T.C., 60–61, 144–145, 176, 186–187, 197 Saiki, K., 337 Saiki, R., 311 Saito, A., 2–4, 7, 11, 14 Saito, T., 60–61, 144–145 Sakai, J., 2, 128–130, 230, 296 Saleh, O., 310, 340–341 Salema, V., 63–64 Salloway, S., 186–187 Salmon, S.E., 49 Sambrook, J., 105 Samuels, M., 313–314 Samuelson, J.C., 45 Sanders, C., 216–217 Sane, D.C., 360–361 Sanjana, N.E., 371–372, 382 Sannerud, R., 84 Sano, Y., 143–144, 158–159 Santoro, M.M., 313–314 Sardana, M.K., 128–129, 297, 399–400 Sargent, F., 45

458 Sasaki, K., 311 Sasaki, T., 189–191, 193–194 Sato, C., 138, 189–198, 192f, 195f Sato, F., 311 Sato, M., 2–4, 22–25 Sato, T., 137, 189–191, 193–194 Sauer, R.T., 2–4, 25 Sauguet, L., 61 Saulnier, J., 420–421 Savidge, B., 314 Sawada, N., 313–314, 316 Schaap, O., 313–314 Schafer, N.P., 100–101, 231 Sch€afer, W., 80–81 Schagger, H., 404 Schall, C., 340–341 Scharnagl, C., 8, 128, 285 Scheek, S., 129–130 Scheer, M., 379 Scheich, C., 138 Scheid, J.F., 78 Scheltens, P., 186–187 Schepens, B., 87 Schepers, U., 130 Scheres, S.H., 61, 148, 189, 193–194 Scherman, M.S., 314, 316 Schettgen, T., 357 Schledz, M., 314, 316 Schleeger, S., 130 Schlegel, S., 102–104, 256–257 Schleif, R., 259 Schliep, J.E., 84–85 Schlieper, D., 396 Schmid, H.C., 315 Schmidt, B., 159 Schnaitman, C.A., 246 Schneider, A., 159–160 Schneider, B., 101–102, 138 Schnoes, H.K., 355–356 Schoch, S., 315–316 Schoolmeester, A., 87 Schoonjans, L., 197–198 Schoonooghe, S., 85 Schr€ oder, B., 130, 296–297 Schroeder, F., 159–160 Schroter, A., 412–414, 428–430 Schuenemann, T.A., 45 Schulman, S., 352–353, 368–369

Author Index

Schultz, G., 314 Schultz, P.G., 27–29 Schulz, S., 315–316 Schulze, D., 323–324, 334, 337 Schulze, E., 337 Schurgers, L.J., 350–351, 357, 386 Schurig-Briccio, L.A., 311–313, 319 Schwaiger, M., 80–81 Schwarz, D., 101–102 Scott, D.A., 382 Scully, M., 87 Seal, R.P., 189–191 Seamone, C., 313–314 Seddon, A.M., 264–265 Seeger, K., 340–341 Seegmiller, A.C., 129–130 Seidel, R., 340 Seidler, A., 314, 316 Sela, M., 80–81 Selkoe, D.J., 8, 128–129, 138, 158–159, 186–189, 230, 240, 296–307 Sepp€al€a, S., 63–64 Sepulveda, J., 313–314 Sepulveda-Falla, D., 60–61, 159–160 Serneels, L., 68–69, 197–198 Seubert, P., 160 Severin, K., 311 Seybold, M., 100–101, 112–114, 285–286 Shah, D.V., 355 Shalem, O., 382 Sharifzadeh, Z., 80–81 Sharpe, H.J., 100, 117–118, 210–214, 256–257, 282–285, 288–290, 396–398, 400–401, 408–409, 421 Shashilov, V.A., 210 Shearer, M.J., 350, 377–378, 386 Shen, L.M., 46–47 Sherratt, A.R., 100–101, 112–114 Sherrington, R., 128–129, 186–187, 208–209 Shewmaker, C.K., 314 Shi, L., 189–191 Shi, X., 382 Shi, Y., 2, 61, 100, 128–149, 142f, 146f, 158, 189, 193–194, 280–282, 397–400 Shiba, S., 379 Shibata, F., 197–198 Shibuya, K., 314 Shimada, N., 159, 175, 189–191, 194–195

Author Index

Shimomura, Y., 313–314, 316 Shineberg, B., 311–313 Shinoda, T., 175, 188–191, 194–196 Shinozaki, K., 176, 186–187, 197 Shirotani, K., 159, 162, 167, 171, 175–176 Shoemaker, C.B., 87, 356–357 Shokhen, M., 280 Shokrgozar, M.A., 80–81 Shukla, A.K., 101–102 Siddiq, S., 377–378 Sieber, S., 285–288, 287f, 397 Silence, K., 87 Silengo, L., 313–314 Silhavy, T.J., 397–398 Silverman, R.B., 352–353 Simpson, J.V., 209–210, 224 Sinning, I., 101–102 Siricilla, S., 311–313 Sisodia, S.S., 208–209 Skerle, J., 100–101, 145–146, 281f, 282–285, 283f, 287f, 288–290, 396 Sˇkerlova´, J., 83 Skiba, P.M., 100, 257–259, 268, 396–397, 400–401, 419–420 Slabbaert, J.R., 313–314 Slayden, R.A., 311–313 Sligar, S.G., 65 Slotboom, D.J., 258 Slunt, H.H., 195–196 Smialowska, A., 167 Smider, V.V., 61–62 Smidt, I., 86–87 Smirnov, A., 144–145 Smit, M.J., 65–66, 86–87 Smith, P.A., 39–40, 45 Smith, P.K., 402 Smith, P.M., 316 Smitha Rao, C.V., 38–39 Smitka, T.A., 39 Sobhanifar, S., 138 Søderberg, J.N., 80–81 S€ oding, J., 316–319 Soll, J., 314 Sommer, S., 311 Song, J., 258–260 Song, Y., 420–421, 431–434 Sonoda, Y., 328–329, 333–334 Sontag, B., 311

459 Sookrung, N., 84 Sørensen, T.L.-M., 101–102 Sornkom, J., 87–88 Soto, M.C., 130 Sottrup-Jensen, L., 355 Soumailakakis, D., 138 Soute, B.A., 352–357, 359–361 Spangler, J.B., 80–81 Spencer, J.S., 314, 337 Sperling, R., 186–187 Spiro, T.G., 209–210, 223 Spitzer, L., 195–196 Spohn, G., 358–359 Sponer, B., 84 Spriestersbach, A., 107–110 Srimanote, P., 84 St George-Hyslop, P.H., 61, 128–129, 137–138, 208–209 Stafford, D.W., 350–387 Sta˚hl, S., 63–64 Stainier, D.Y.R., 313–314 Stam, J.C., 87–88 Stanchev, S., 100–101, 145–146, 281f, 282–285, 283f, 287f, 288–290, 396 Standage, D.S., 372 Stanley, E.C., 71 Staufenbiel, M., 144–145 Staus, D.P., 85–86 Stec, E., 337, 340–341 Steed, D.B., 39 Steeland, S., 81 Stehle, T., 340–341 Steidl, S., 83 Stein, R.L., 43, 47–48 Steinaa, L., 80–81 Steiner, H., 8, 120–121, 128, 137–138, 143, 158–181, 187–188, 240, 285, 296–297, 303–304 Steitz, T.A., 208–209 Stenflo, J., 350–351, 355 Stern, E.A., 60–61 Stevens, R.D., 378–379 Stevenson, C.E.M., 340–341 Stevenson, D., 311–313, 316 Stevenson, L.G., 208–209, 239–240, 282–284, 400, 419–420 Steyaert, J., 63–64, 66–67, 71–72, 80, 84–86 Stigter-van Walsum, M., 65–66, 87–88

460 Stohr, T., 86–87 Stone, M.O., 70 Stortelers, C., 65–66 Story, B., 350–351 Strachan, R.T., 85–86 Straight, D.L., 358–361, 364–365, 367–368, 378–379, 381–382 Strege, M.A., 39 Strisovsky, K., 100–101, 112–114, 117–118, 145–146, 210–214, 239–240, 248, 256–274, 279–290, 281f, 283f, 287f, 396–433 Strohner, R., 83 Strokappe, N.M., 86–87 Strub, J.-M., 298 Struhl, G., 158 Strynadka, N.C., 36–41, 43, 47, 280–282 Sturgill, G.M., 419–420 Suciu, D., 41, 43 Suen, E.T., 355–356 Suhara, Y., 313–314, 316 Suharni, 65 Sui, S.F., 258–260 Sulli, C., 84 Sun, H., 311–313, 319, 379 Sun, J., 81 Sun, L., 2, 100–102, 142f, 144–145, 146f, 147, 158, 189, 191–193, 297 Sun, P., 84 Sun, P.M., 36 Sun, S., 83 Sun, T., 81 Sun, Y.M., 360–361 Sungsuwan, S., 100–101 Suttie, J.W., 352–353, 355–361 Suvarna, K., 311–313, 316 Suzuki, K., 311 Suzuki, M., 2–4 Suzuki, N., 186–187 Svendsen, I., 50–51 Svensson, L., 86–87 Svergun, D.I., 83 Swain, R.K., 285–286 Swanson, R.V., 340 Sweeney, M.C., 50–52 Swiezewska, E., 311, 316 Sym, M., 187–188 Szaruga, M., 60–61, 159–160

Author Index

T Tabata, S., 2–4, 22–25 Tachiya, M., 233–235 Tagami, S., 158–159 Takagi, S., 189–193, 192f, 195f, 197 Takagi-Niidome, S., 189–191, 193–197 Takahashi, Y., 137–138, 187–188, 194, 296–297, 314 Takai, Y., 320, 401, 421 Takami, M., 143–144, 158–159 Takashima, M., 78 Takasugi, N., 137–138, 187–188, 197, 296–297 Takeda, M., 137 Takegami, N., 175, 188–191, 194–196 Takeo, K., 175, 188–191, 194–196 Takeuchi, H., 210 Takeuchi, S., 361–362 Takikawa, R., 197 Tamura-Kawakami, K., 2–4, 22–25 Tanaka, K., 311 Tandon, A., 187–188 Tanimura, S., 175, 188–191, 194–196 Tanimura, Y., 158–159, 188 Taouji, S., 78 Tarcz, S., 340–341 Tari, L.W., 340 Tarry, M., 102–104, 256–257 Tate, C.G., 414–415 Taussig, M.J., 63–64 Taverna, M., 362–363, 378 Tchouate Gainkam, L.O., 87–88 Temperton, N.J., 87 Tersago, D., 87 Teunissen, K.J., 386 Thakkar, A., 50–52 Thanongsaksrikul, J., 84 Theuns, J., 196–197 Theuwissen, E., 386 Thijssen, H.H., 352–354, 359–360 Thinakaran, G., 195–196 Thompson, M.K., 78 Thueng-in, K., 84 Tie, J.K., 350–387 Tie, K., 358–359, 372, 374, 377–378 Ting, Y.T., 36, 38 Tokuda, H., 14, 17 Tokuhiro, S., 176, 186–187, 197

461

Author Index

Tokunaga, F., 361–362 Tol, M.B., 61–62 Tolia, A., 79–80, 189–191 Tomaszewski, J., 313–314 Tominaga, A., 189–197 Tominari, Y., 194 Tomioka, M., 159–160, 162, 167, 171 Tomita, M., 320 Tomita, T., 137–138, 176, 186–201, 192f, 195f, 296–297 Toomey, J.R., 361–362 Torres-Arancivia, C., 136–137, 208–209, 211–216 Townsend, M.G., 353–354 Trakhanov, S., 84–85 Trambauer, J., 158–181, 240, 303–304 Trankle, C., 412–414, 428–430 Trauger, S.A., 45 Trelinski, J., 80–81 Tremblay, J.M., 87 Triest, S., 64, 66–67, 71–72, 80 Tripathy, A., 357–358 Tromp, G., 313–314 Tropea, J.E., 272–274 Truong, H.H., 100 Truusalu, K., 86–87 Tsai, C.J., 61, 193–194 Tsai, J.Y., 186–187, 198, 289–290 Tschantz, W.R., 36, 40, 43, 45 Tsubaki, M., 337 Tsukui, T., 379 Tsuruoka, M., 137–138, 187–188, 296–297 Turbyfill, J.L., 358–359 Turco, E., 313–314 Turcotte, S., 78 Turner, S.H., 366 Tyndall, J.D., 282–284 Tyree, C., 159–160 Tzagoloff, A., 316

U Ubarretxena-Belandia, I., 136–137, 208–225, 258 Uchida, N., 311 Uchino, Y., 313–314, 316 Ueda, M., 311 Uemura, K., 60–61, 159, 175, 189–191, 194–195

Ulrich, M.M., 356–357 Ulrichts, H., 87 Umezawa, N., 194 Unger, M., 87–88 Uns€ old, I., 340–341 Upadhyay, A., 311–313, 319 Urano, Y., 65–66 Urban, S., 2, 8, 100–101, 121, 128–129, 135–136, 145–146, 208–209, 211–212, 230–250, 256–258, 265–266, 279–280, 282–286, 287f, 288–290, 296–297, 299–305, 396–400, 408–409 Uritsky, N., 280 Usubalieva, A., 359–361

V Vaia, R.A., 70 Valentin, H.E., 314 van Belzen, I.A., 283f van Bergen En Henegouwen, P.M.P., 87–88 Van Blarcom, T.J., 81 Van Broeck, B., 196–197 van den Brink-van der Laan, E., 39–40 Van Den Broeck, T., 85 van Dijl, J.M., 36 van Dongen, G.A.M.S., 87–88 Van Doren, S.R., 430–431 Van Gassen, N., 75 van Haarlem, M., 352–354, 360–361 Van Hauwermeiren, F., 81 Van Heeke, G., 86 van Klompenburg, W., 39–40, 43 van Lare, J., 186–187 Van Leuven, F., 137 Van Meensel, S., 313–314 Van Solt, C.B., 86–87 Van Valckenborgh, E., 87–88 Van Zijderveld, F.G., 86–87 Van Zijderveld-Van Bemmel, A.M., 86–87 Vandenbroucke, K., 86–87 Vandenbroucke, R.E., 81 Vanderkerken, K., 87–88 Vandersteen, A., 60–61, 144–145, 188, 196–197, 298 Vanderstichele, H., 137 Vandyk, J.K., 372 Vaneycken, I., 75 Vanhove, C., 87–88

462 Vanlandschoot, P., 65–66 Vanstreels, E., 84–86 Vardanian, L., 163 Vasina, J.A., 259–260 Vasylieva, N., 87 Veenstra, A., 419–420 Venema, G., 36 Venkatesh, T.V., 314 Venken, T., 84 Vennekens, K., 196–197 Vercruysse, T., 84–86 Vereecke, D., 314, 316, 337 Veres, T., 85 Verheesen, P., 64–66, 70–71 Verhelst, S., 145–146, 248, 257–258, 283f, 285–286, 287f, 396 Verhelst, S.H., 256–257, 285–289, 287f, 396–397 Verhelst, S.H.L., 100–101, 112–114 Verkaar, F., 86 Vermeer, C., 350–357, 360–361, 383–386 Verstreken, P., 313–314 Vervoort, L.M., 359–360 Vervoort, L.T., 352–353 Veugelen, S., 60–88, 144–145, 159–160, 193–194 Vibat, C.R.T., 63–64 Vilain, S., 313–314 Villa, A., 259–260 Vincke, C., 75, 81, 87–88 Vinothkumar, K.R., 100–101, 145–146, 248, 257–258, 283f, 285–289, 287f, 396 Voet, A., 84 Volkov, Y., 85 von Depka Prondzinski, M., 80–81 von Heijne, G., 39–40, 43, 45–47, 333–334 Vos, M., 313–314 Voss, M., 130, 296–297 Vosyka, O., 100–101, 285–289, 287f, 396–397 Voytas, D.F., 372 Vrentas, C., 87 Vreugde, S., 130

W Wackernagel, W., 322 Wadelius, M., 370 Wagner, S., 102–104, 256–257

Author Index

Wa˚hlin, L., 86 Wajih, N., 352–353, 360–361 Wakabayashi, S., 361–362 Walker, B., 289 Walker, J.E., 102–104, 256–257, 326–328 Wallace, B.J., 320–323 Wallach, J., 420–421 Wallin, R., 352–354, 358–361 Wallrapp, F.H., 340 Walls, J.D., 361 Walter, J., 176 Wan, B., 311–313 Wang da, N., 258–259 Wang, B., 352–353, 368–369 Wang, D.N., 258–260, 266–267, 414–415 Wang, F., 101–102 Wang, H., 313–314 Wang, J., 132–133, 136–138, 146f, 147 Wang, L., 36 Wang, P., 36, 39–41, 46 Wang, Q.M., 36 Wang, R., 159–160, 189, 298 Wang, X., 368–369 Wang, Y., 2, 100–101, 116, 131, 135–136, 145–146, 208–210, 221–222, 231, 257–258, 265, 280–282, 297, 396–397, 409 Wang, Z., 2, 131, 136, 146, 146f, 297 Wanner, B.L., 320 Ward, J.M., 71 Wasserman, J.D., 128 Wasson, P., 313–314 Watanabe, M., 313–314, 316 Watanabe, N., 189–191, 194–198 Watzka, M., 358–360, 362–363, 378–379 Waugh, D.S., 272–274 Wavreille, A.S., 50–52 Weber, R., 337 Weggen, S., 158–160 Wei, J., 350–351 Weihofen, A., 2, 128–130, 187–188, 208–209, 230, 297 Weinfurtner, D., 83 Weinglass, A.B., 189–191 Weis, W.I., 61 Weiss, J.D., 314 Weiss, J.S., 313–314 Wendel, S., 63–64

463

Author Index

Wenthur, C.J., 61 Wernery, U., 63–64, 75 Wessjohann, L.A., 311, 313–314, 323–324, 334, 337 Westhofen, P., 359–360 Westwood, O.M.R., 83 White, P.S., 313–314 Whitlon, D.S., 353, 358–359 Wickner, W., 36, 44–45 Wiegering, V., 379 Wieland, R., 379 Wijetilaka, R., 398 Willems, B.A., 350–351 Williams, H., 39–40 Williams, J., 353–354, 359–360 Willows, R.D., 315 Wilson, C.J., 39–40 Wilson, L.A., 359–361 Wiltfang, J., 120, 144–145 Windyga, J., 80–81 Winge, D.R., 316 Wingler, L.M., 85–86 Winkler, E., 137–138, 171, 176, 187–188, 240, 296–297, 303–304 Winter, G., 70 Winters, R.S., 313–314 Wittenburg, N., 163 Wittrup, K.D., 80–81 Wohlk€ onig, A., 64, 66–67, 71–72, 80 W€ ohri, A.B., 336 Wolf, E.V., 100–101, 112–114, 256–257, 285–289, 287f, 396–397 Wolfe, M.S., 2, 8, 36, 60–61, 100–101, 128–129, 135, 137–138, 143–145, 158, 187–189, 208–209, 211–212, 230–231, 240, 248, 256–258, 265–266, 289–290, 296–307, 396–400 Wolfe, P.B., 36 Wolynes, P.G., 100 Wong, P.C., 79–80 Wong, Y.-H.H., 314 Woodcock, J., 370 Wright, D.J., 378–379 Wu, H., 87 Wu, H.L., 285–286 Wu, J., 189–191, 272–274 Wu, Q., 223 Wu, S., 136–137

Wu, S.M., 356–357 Wu, Z., 2, 130–131, 135–136, 145–146, 146f, 208–209, 231, 280–282, 297 Wunderlich, F.T., 358–359 Wyss, R., 61–62

X Xavier, C., 75, 87–88 Xia, W., 128–129, 187–188, 230, 296 Xie, T., 101–102, 142f, 144, 189 Xie, X., 340–341 Xu, M., 128–129, 186–187, 194, 296–297, 399–400 Xu, Y., 81 Xu, Z.L., 87 Xue, Y., 100–101, 112–114, 145–146, 285–286, 287f, 409

Y Yager, D., 159–160 Yakubenko, A.V., 360–361 Yamamoto, H., 311 Yamamoto, K., 311 Yamane, H., 311 Yamasaki, A., 137–138, 167, 176 Yan, C., 2, 101–102, 132–133, 136–138, 142f, 144–145, 146f, 147, 189, 191–193, 297 Yan, H., 2, 130–131, 135–136, 145–146, 146f, 231, 280–282, 297 Yan, N., 2, 130–131, 135–136, 145–146, 146f, 231, 280–282, 297 Yan, S.C., 361 Yanagida, K., 158–159 Yang, G., 2, 61, 128–149, 146f, 189, 191–194, 297 Yang, H., 311–314, 319 Yang, L., 380–381 Yang, Y., 310–341 Yang, Z., 100–101 Yankner, B.A., 128 Yao, X.J., 61 Yarden, Y., 80–81 Yashiro, S., 333–334 Yau, Y.H., 86 Yazaki, K., 311, 337 Ye, J., 2, 36, 128–130, 230, 296 Ye, W., 138, 187–189, 296–300

464 Yeung, Y.A., 81 Yin, J.A., 27–29 Yokoshima, S., 194 Young, I.G., 311–313, 316, 320–323 Young, K., 100, 135–136, 397–400 Young, P.G., 36, 38 Young, T.S., 27–29 Yu, G., 187–188 Yu, H., 83 Yuasa, M., 311 Yuge, N., 313–314, 316

Z Zahariev, I.K., 175, 188–191, 194–196 Zahnd, C., 63–64 Zakharova, S., 323–324, 334, 337 Zanaletti, R., 49–50 Zaucke, F., 358–359 Zauner, T., 419 Zeiler, E., 285–288, 287f, 397 Zeissler, A., 256–257, 285–288, 287f, 397 Zelenski, N.G., 2, 128, 230, 296 Zeltins, A., 65–66 Zettl, M., 256–257, 397–398 Zhang, D., 314, 316 Zhang, F., 371–372, 380–382 Zhang, J., 84, 187–188 Zhang, L., 187–188 Zhang, X., 2–4, 159–160 Zhang, Y., 2, 100–101, 116, 131, 145–146, 208–209, 231, 257–258, 265, 280–282, 289, 297, 340, 396–397, 409 Zhang, Z., 128–129

Author Index

Zhao, G., 83 Zhao, X., 84 Zhao, Y., 86–87 Zhao, Y.J., 87–88 Zhen, J., 258–259 Zheng, F., 85 Zheng, H., 143 Zheng, S., 148 Zhou, L., 84 Zhou, M., 315–316, 319, 330, 334, 337 Zhou, R., 128–149 Zhou, Y., 282, 289, 371–372 Zhou, Y.-H., 83–84 Zhou, Z., 135, 258–259 Zhu, A., 379 Zhu, H.Y., 45 Zhu, L., 258 Zhu, W., 311–313, 319 Zhu, X., 311 Zimmer, A.K., 137 Zimmer, J., 330 Zimmermann, A., 352–353 Zineh, I., 370 Zinn, K., 16 Zocher, G., 340–341 Zoll, S., 100–101, 145–146, 281f, 282–285, 283f, 287f, 288–290, 396 Zolnerciks, J.K., 65 Zornig, M., 130 Zuchner, T., 419 Zuckermann, R.N., 50–52 Zwizinski, C., 36 Zytkovicz, T.H., 350–351

SUBJECT INDEX Note: Page numbers followed by “f ” indicate figures, and “t” indicate tables.

A Ablynx, 86–87 4-Acetamido-40 -maleimidylstilbene-2, 20 -disulfonic acid (AMS) to Cys residues, 18–20, 19f modification assay, 20–22 Activity-based probes (ABPs), 285–288 Activity-based profiling, 100–101, 112 AICD. See APP intracellular domain (AICD) AlphaScreen, 72–75, 73f, 78 Alzheimer’s disease (AD), 158 AD-linked PSENs, 60–61 amyloid precursor protein in, 296–297 β-amyloid peptides, 208–209 γ-secretase, 186–187 Amyloid-β (Aβ), 158–159, 186–187 immunoprecipitation, 171 mass spectrometry analysis, 172–175 MSD sandwich immunoassay analysis, 162–166 Tris–Bicine urea SDS-PAGE analysis, 166–172 Amyloid precursor protein (APP), 128–129, 143, 186–188, 186f, 189f, 296–297 Antigen elution methods, 70–71 γ-secretase, methods to capture, 67–70 presentation on solid phase, 66–67 in solution approach, 67 sources, 65–66 Anti-γ-secretase nanobodies, 62 AlphaScreen, 73–75 characterization, 82f A. pernix UbiA (ApUbiA), 330–331 APP. See Amyloid precursor protein (APP) APP intracellular domain (AICD), 143–145 D-Arginine, 47–48 AV12 cells, 359, 361–362, 367–368

B Bacteria cytoplasmic membrane, 14

pathogenic, drug resistance of, 38–39 rhomboid proteases, 101–112 S2P family protease, 3f survival role, 2–4 in vivo assays, 397–398 Beta-lactams, 286–289 Beta-lactones, 286–288 Bicinchoninic acid (BCA) assay, 272, 274, 402, 422 Biopannings antigen elution methods, 70–71 γ-secretase, methods to capture, 67–70 on solid phase, 66–67 in solution, 67 sources, 65–66 critical aspects of, 64–71 whole cells and virus-like particles, 65–66 Biotinylation γ-secretase, 67–68, 194–195 intact cell, 198–199 SCAM using microsome, 199–200 β-lactamase (Bla), 7, 12f BODIPY dye, 409–418 BODIPY FL dye, 115, 116f

C CBB. See Coomassie Brilliant Blue (CBB) CD. See Circular dichroism (CD) cDNA GGCX, 383 prothrombin, 381–382 VKOR, 375 Cell-based screening assays recombinant protein, 75–77 vitamin K cycle enzymes, 361–369, 362f Cell culture GSMs, 161–162 medium, 361–365, 375 and protein expression, 105 465

466 Cell lysis high-pressure homogenizer method, 264–265 and membrane fraction isolation, 267–268 Cholesteryl hemisuccinate (CHS), 107–110 Circular dichroism (CD), 305 ClonePix FL technology, 364 Cloning, UbiA superfamily, 325–326 Combinatorial peptide library, 38, 48f, 51f Coomassie Brilliant Blue (CBB), 8, 13 Copper-phenanthroline, cross-linking experiment, 201 COQ2, 311, 319, 339 CRISPR-Cas9-mediated gene knockout reporter cells, 378–386 Cryoelectron microscopy (cryo-EM), 129, 148 Crystallization in detergents, 334–335 in LCP, 335–337 Cysteine-based structural analyses, presenilin cross-linking experiment, 195–196, 196f γ-secretase, biotinylation of, 194–195 genetic mutation effect, 196–197 SCAM, 189–191, 190f, 192f topology and position of Cys, 191–194, 193f Cysteine (Cys) residue AMS to, 18–20, 19f mutant presenilin 1 expressing cell, 197–198 topology and position of, 191–194, 193f

D Deep-UV resonance Raman (dUVRR) spectroscopy, 208–209, 222f intramembrane proteolysis, 209–210, 222f measurements, 210–211, 219 spectral acquisition and analysis, 223–224 Delineation of epitope, 83–84 Detergents crystallization in, 334–335 FL-casein, 412–413 GlpG activity, 412–413, 414f intramembrane proteolysis in, 219–220 kinetic analysis of proteolysis, 233–235

Subject Index

psTatA-FRET, 428–429, 429–430f selection of, 333–334 steady-state kinetic analysis in, 234f Detergent-solubilized assay, 297–299 Disulfide cross-linking, 29–31, 31f Dithiothreitol (DTT), 352–354, 370 DNA ethanol precipitation, 119 Double-gene knockout (DGKO) reporter cells, 374, 374f VKOR using, 375–376, 377f Drosophila Rhomboid-4, 100, 235 dUVRR spectroscopy. See Deep-UV resonance Raman (dUVRR) spectroscopy

E Electrophiles, 285–289 ELISA. See Enzyme-linked immunosorbent assay (ELISA) Elution methods, 70–71 EnzChek protease assay, 115–117 Enzymatic activity γ-secretase, 143–145 HPLC analysis, 325 intramembrane prenyltransferases, 319–325 intramembrane proteases, 135–137 in microsomes, 323–325 Enzyme-linked immunosorbent assay (ELISA), 361–362, 362f, 364–366, 367f in-cell, 75–77, 76f recombinant protein, 72 Epitope, 80–84 binning method, 81–83 delineation, 83–84 Escherichia coli (E. coli) BODIPY FL dye, 115, 116f GlpG, 103f, 108f, 113f, 116, 116f rhomboid expression and purification workflow, 103f rhomboid protease, 280 RseP (see RseP) signal peptidase, 36, 38–39

F FIXgla-PC reporter cell lines, 363–364, 365f, 367f

467

Subject Index

FL-casein. See Fluorescently labeled casein (FL-casein) Flow cytometry, 75 Fluorescence quenching, inducible proteoliposome assay, 305 Fluorescence resonance energy transfer (FRET) peptide library method, 47, 48f, 50–51, 51f resin-bound substrate library approach, 53 wild-type and SP mutants with, 53t Fluorescence size exclusion chromatography (FSEC), 329 Fluorescently labeled casein (FL-casein), 409–418 concentration, 415–416, 416f GlpG-mediated cleavage, 411–412, 416–417, 417f Michaelis–Menten kinetics, 416–417, 419f optimal detergent concentration, 412–413 time course, 411f Fluorescent substrates, 408–433 BODIPY-casein, 409–418 FRET-psTatA, 419–433 Fluorophore-labeled substrate, 239 Forster resonance energy transfer (FRET), P. stuartii TatA, 419–421, 420f AarA-mediated cleavage, 429f, 431–433 expression and purification, 421–423 fluorescence and concentration, 430–431, 431f kinetic assay parameters, 423–424 Michaelis–Menten kinetics, 424, 433f optimal detergent concentration, 428–429, 430f optimal rhomboid concentration, 424–426 pH activity, 426–427, 427f reliability, 423 temperature, 427–428, 429f time course, 425f FP-labeled rhomboids, 112–114 French press, 264–265 Functional-based screening assay, γ-secretase, 78–80, 79f

G Gamma-glutamyl carboxylase (GGCX) activity assays, 357 artificial peptide substrates, 356 cDNA, 383 CRISPR-Cas9-mediated gene knockout reporter cells, 378–386, 381f function study, 355–358, 358f genetic screening for, 379 GGCX-deficient reporter cells, 383–384 vitamin K titration, 385f γ-secretase, 137–138, 158, 297, 303 amide coupling, 69–70 biotinylation, 67–68, 194–195 CHAPSO-solubilized assay, 298–299 complexes, 60 enzymatic activity assay, 143–145 functional-based screening assay, 78–80, 79f immunocoupling, 68–69 in-cell ELISA, 75–77, 76f intramembrane cleavage, 188 mammalian expression vector pMLink, 138–140, 139f molecular basis of, 187–188 pMLink vector, 138–140, 139f purification, 141–143, 142f on sepharose beads, 69–70 transient expression, 140 γ-secretase inhibitors (GSIs), 159, 186–187, 194–195, 195f γ-secretase modulators (GSMs), 159–160, 160f acidic and nonacidic, 165f amyloid-β mass spectrometry analysis, 172–175 MSD sandwich immunoassay analysis, 162–166 Tris–Bicine urea SDS-PAGE analysis, 166–172 assaying activity, 160–175, 165f cell culture and drug treatments, 161–162 photoaffinity labeling, 175–181, 175f Genetic screening, 129 GlpG activity, 396–397 dependence on pH, 413f detergent concentration, 231, 414f E. coli, 103f, 108f, 113f, 116, 116f

468 GlpG activity (Continued ) expression, 130–131 purification, 133–135 SDS-PAGE of FL-casein cleavage, 417f SEC on, 414–415 solubilization conditions, 110 G protein-coupled receptors (GPCRs), 61 Green fluorescent protein (GFP), 258, 329, 399–400, 419

H HEK293 cells, 358–359, 361, 366, 379–382 vitamin K reduction in, 367–368 VKOR, 373, 376–377 High-pressure homogenizer, 264–265 High-throughput sequencing (HTS), 78 His-tagged proteins, 268–269, 272–274 HPLC analysis, enzymatic activity, 325 HPLC-based VKOR, 353–354, 354f Hydrolytic process, 100

I I-CLiPs. See Intramembrane-cleaving proteases (I-CLiPs) IMAC. See Ion metal affinity chromatography (IMAC) Immunocoupling, γ-secretase, 68–69 Immunoprecipitation amyloid-β, 167, 171 γ-secretase, 81 IMVs. See Inverted membrane vesicles (IMVs) In-cell ELISA, 75–77, 76f Inducible proteoliposome assay, 301–306 flow chart, 302f methods, 304 verification of enzyme and substrate, 305–306 Inhibitors γ-secretase inhibitors, 159 rhomboid protease, 287f first generation, 285–288 future perspectives, 288–290 structure-guided design, 288–290 serine protease, 285–286 Intact cell biotinylation, 198–199 SCAM using, 198–199

Subject Index

Intramembrane-cleaving proteases (I-CLiPs), 2, 296–297 active site, 18–20 detergent-solubilized assay, 297–299 enzymatic assay, 136f expression and purification, 134f inducible proteoliposome assay, 301–306 involvement, 129 RseP–substrate interaction, 27 structures, 145–147, 146f Intramembrane prenyltransferases crystal structures of, 338–339f sequence clustering analysis, 315–318, 317f soluble vs., 337–341, 338–339f UbiA superfamily of (see UbiA superfamily) Intramembrane proteases, 128–129, 208–209, 279–280 activity assay, 212f enzymatic activity assays, 135–137 high-resolution structures of, 100 inducible and fluorogenic assay, 232f, 237f characterization of experimental system, 245 extended progress curve analysis, 243–244 gel analysis, 242–243 substrate and, 240–242, 244–245 kinetic analysis in detergent micelle systems, 233–235 inhibition, 248–250, 249f purification procedure, 216–218 substrates, 282–284 transmembrane domain, 208–209 ubiquity of, 230–231 Intramembrane proteolysis in detergent, 219–220 dUVRR spectroscopy, 221–224, 222f substrates purification, procedure for, 214–216 in vitro assay, 211–221 Inverted membrane vesicles (IMVs), 11, 14 preparation and in vitro translocation, 14–16, 15f RseP, 17–18

Subject Index

In vitro assay GGCX, 355–358, 358f intramembrane proteolysis, 211–221 rhomboid proteases, 398–399 fluorescent substrates, 408–433 gel-based, 399–408 transmembrane protein substrates, 399–408 VKOR, 352–354, 354f VKR, 358–361 In vitro signal peptide cleavage assay, 43–44 In vivo assay mal-PEG modification, 23f, 25–27 rhomboid proteases in bacteria, 397–398 in mammalian cells, 398 RseP proteolytic activity, MBP, 5–8, 6f In vivo cross-linking analysis, 27–31 In vivo imaging, nanobodies for, 87–88 Ion metal affinity chromatography (IMAC), 268–272 Isocoumarins, 286–288 Isotopically labeled enzyme, 219

K Kanamycin-resistant (KanR) gene, 320, 322

L Leader peptidase (Lep), 3f, 7 arabinose-dependent strain, 45 Liposomes preparation of, 238–239 unilamellar, 238–239 Liquid cubic phase (LCP), 335–337

M MALDI-MS analysis, 47–48, 51–52 mal-PEG modification, 23f, 25–27 Maltose-binding protein (MBP), 43 RseP proteolytic activity, 4–11, 6f Mammalian cells, in vivo assays, 398 Mammalian expression vector, pMLink, 138–140, 139f Mass spectrometry (MS) analysis, 220–221 amyloid-β, 172–175

469 Matrix-assistedlaserdesorption/ionization (MALDI)-time of flight (TOF) mass spectrometry (MS), 166, 220–221 Matrix Gla protein (MGP), 350–351, 381–382, 386 carboxylation defects, 379–380 domain structure, 380f moderate effect on, 385–386 MBP. See Maltose-binding protein (MBP) Membrane-associated proteases, 2 Membrane fraction isolation, 267–268 solubilization, 270–272 Membrane-immersed proteolysis intramembrane protease inhibition assay, 248–250, 249f liposomes preparation, 238–239 real-time kinetic analysis, 245–250, 248f steady-state kinetic analysis, 245–248 Membrane protein biochemical analysis of, 235–236 biogenesis pathway, 256 crystallography, 315, 328–329 extraction, 266 pBAD expression system, 259 production, 263–265 purification, 257–258 recombinant, 265 Menaquinones, 310, 320, 386 Meso Scale Discovery (MSD), 162–166 Metalloproteases, 208–209 site-2 protease family, 2–4, 230 Methanethiosulfonate (MTS), 189–193, 201 Methanocaldococcus jannaschii, 131, 146, 146f [35S]Met-labeled model presecretory protein, 17–18 MGP. See Matrix Gla protein (MGP) Michaelis–Menten kinetics fluorescently labeled casein, 416–417, 419f psTatA-FRET, 424, 433f Microsomes, 360–361, 370 enzymatic activity in, 323–325 purified, 352 SCAM using, 199–200 wild-type EcUbiA in, 326f mRNA preparation, 16–17

470 MS analysis. See Mass spectrometry (MS) analysis MSD. See Meso Scale Discovery (MSD)

N Nanobodies, 62 affinity determination, 84–85 applications, 63f, 85–88 biopannings, 64–71 characterization, 63f clinical and industrial applications, 86–87 conformation-sensitive, 62 crystallization of proteins, 86 epitope, 80–84 functional screenings, 78–80 fundamental biochemical and cellular studies, 85–86 high-throughput sequencing, 78 identification, 63f phage display, 62–64 screening for binders, 71–80 AlphaScreen, 72–75, 73f cell-based screening assays, 75–77 enzyme-linked immunosorbent assay, 72 recombinant protein, 72–75 sequencing-based methods, 77–78 for in vivo imaging, 87–88 Nebulization, 86–87 N-hydroxysuccinimide (NHS) beads, γ-secretase, 69–70 Nonhomologous end-joining (NHEJ) pathway, 371–372, 371f, 380–381 Nonsteroidal antiinflammatory drugs (NSAIDs), 159–160

O On-bead screening, of peptide library, 50–52 Oxyanion pocket, 282

P Parkinson’s disease, 230 Partial Edman degradation–mass spectrometry (PED-MS), 47–48, 50–53, 51f pBAD system, rhomboid proteases, 258–274 Peptide library combinatorial, 38, 48f, 51f

Subject Index

design, 47–48 on-bead screening of, 50–52 reagents, 48 screening, 52 synthesis, 49–50 Peptide sequence, 53 Periplasmic domain, RseP, 22–25, 23f Phage display, 62–64 Photoaffinity labeling, GSMs using, 175–181, 175f pMLink vector, 138–140, 139f Polyvalent display, 62 pONA. See ProOmpA nuclease A (pONA) Presenilin (PS), 128–129 Cys mutants, 197–198 cysteine-based structural analyses cross-linking experiment, 195–196, 196f γ-secretase, biotinylation of, 194–195 genetic mutation effect, 196–197 SCAM, 189–191, 190f, 192f topology and position of Cys, 191–194, 193f deposition of, 186f FAD-linked mutations on PS1, 186f, 188–191, 196–197 interacts with novel proteins, 187–188 structural characteristics, 195–196 transient expression, 197–198 Presenilin/SPP from archaebacterium (PSH) glimpse, 129–130 proteolysis assay for, 136–137 purification, 132f, 133–135 sequence-based protein engineering, 131–133, 132f ProOmpA nuclease A (pONA), 41–43, 46f Protein Data Bank (PDB), 101 Protein disulfide isomerase (PDI), 352–353 Protein engineering, PSH, 131–133, 132f Proteoliposomes, 235–236, 299–301 inducible, 301–306, 302f quantitative analysis in, 235–238 Proteolytic profiles, determination of, 220–221 Providencia stuartii twin arginine transporter A (psTatA), 239–240, 400 AarA-mediated cleavage, 260–261, 406–408, 407f arabinose concentration, 262f

Subject Index

on expression time, 263–264, 264f temperature, 260f chemiluminescence and concentration, 405, 406f expression and purification, 401–402 FRET, 419–433, 420f AarA-mediated cleavage, 429f, 431–433 expression and purification, 421–423 fluorescence and concentration, 430–431, 431f kinetic assay parameters, 423–424 Michaelis–Menten kinetics, 424, 433f optimal detergent concentration, 428–429, 430f optimal rhomboid concentration, 424–426 pH activity, 426–427, 427f reliability, 423 temperature, 427–428, 429f time course, 425f SDS-PAGE, 402f time course, 404f PS. See Presenilin (PS) Pseudoxanthoma elasticum (PXE)-like syndrome, 379 Purification Δ2-75 SP1, 39–41 GlpG, 133–135 human γ-secretase, 141–143, 142f I-CLiPs, 134f intramembrane proteases, 216–218 membrane protein, 257–258 PSH, 132f, 133–135 rhomboid proteases, 110–112, 257–258, 268–274, 270f RseP, 8–10, 9f S2P, 133–135 SP1 substrate pONA, 41–43 substrates, 214–216 workflow using E. coli, 103f PXE-like syndrome. See Pseudoxanthoma elasticum (PXE)-like syndrome

R Recombinant protein AlphaScreen, 72–75, 73f antigen, 65 enzyme-linked immunosorbent assay, 72 screening on, 72–75

471 Regulated intramembrane proteases, 128 Regulated intramembrane proteolysis, 2, 100 Respiratory syncytial virus (RSV), 86–87 Rhomboid protease, 100, 396–397. See also Intramembrane proteases activity-based probes, 285–288 on arabinose concentration, 262f biological roles of, 279–280 cleavage of chimeric/tagged substrates, 120–123, 121f cleavage products, 213t concentration and reaction time, 403–405 concentration for kinetic analysis, 410–411 detergent screening, 109, 109t EnzChek protease assay, 115–117 Escherichia coli, 280–282 expected timeline, 274 expression, 100–101, 256–257 membrane preparation, 100–107 parameters to adjust, 101–102 fluorescent substrates, 408–433 glimpse of, 129–130 Haemophilus influenzae, 257–259, 269 induction time optimization, 263–264 inhibitors, 285–290, 287f first generation, 285–288 future perspectives, 288–290 structure-guided design, 288–290 ion metal affinity chromatography, 270–272 large-scale expression of, 264–268, 264f large-scale solubilization, 110 mechanism, 280–282, 281f optimal arabinose concentration, 261–263 optimal temperature for, 259–261, 260f parasite-encoded, 230 pBAD system, 258–274 psTatA-FRET, 419–433, 420f AarA-mediated cleavage, 429f, 431–433 expression and purification, 421–423 fluorescence and concentration, 430–431, 431f kinetic assay parameters, 423–424 Michaelis–Menten kinetics, 424, 433f

472 Rhomboid protease (Continued ) optimal detergent concentration, 428–429, 430f optimal rhomboid concentration, 424–426 pH activity, 426–427, 427f reliability, 423 temperature, 427–428, 429f time course, 425f purification, 110–112, 257–258, 268–274, 270f serine hydrolase active site probes, 112–115, 114f size-exclusion chromatography, 258, 272–274, 275f solubilization, 100–101, 107–110, 108f substrate specificity of, 282–285, 283f tag removal, 272–274 transmembrane domain, 117–120, 282–285 in vitro assays, 398–433 fluorescent substrates, 408–433 gel-based, 399–408 transmembrane protein substrates, 399–408 in vitro probing, 112–123 in vivo assays, 397–398 RseP, 2–4 AMS to Cys residues, 18–20, 19f bacterial S2P family protease, 3f cytoplasmic domain, 2–4 disulfide cross-linking, 29–31, 31f mal-PEG modification, 23f, 25–27 membrane domain, 2–4 periplasmic domain, 22–25, 23f proteolytic activity, MBP, 4–11 purification, 8–10, 9f in vivo cleavage assay, 5–8, 6f proteolytic reaction, 10–11 synthetic Bla SP, 12f thiol-specific modification reagents, 18–27 Val13 residue, 20 in vitro analysis of SPs proteolysis, 11–18 in vivo photo cross-linking, 27–29, 28f

S Sandwich immunoassay, MSD, 162–166 SCAM. See Substituted-cysteine accessibility method (SCAM)

Subject Index

SDS-PAGE, 107, 112, 135, 214–216 Bis-Tris SDS-PAGE system, 13 FL-casein cleavage, 417f psTatA purification, 402f Tris–Bicine urea analysis, 166–172 SEC. See Size-exclusion chromatography (SEC) Selenomethionine (SeMet) labeling, 330–331 Sepharose beads, γ-secretase on, 69–70 Sequence clustering, intramembrane prenyltransferases, 315–318, 317f Sequencing-based methods, 77–78 Serine hydrolase active site probes, 112–115, 114f Serine proteases, 36, 280 catalysis, 282 inhibitors, 285–286 Ser-Lys active site architecture, 37–38 Signal peptidases, 36–39 catalytic domain, 38 Escherichia coli, 36, 38–39 purification Δ2-75, 39–41, 44f pONA, 41–43 in vivo assay, 44–46 Signal peptide peptidase (SPP), 128–130, 296–297 Signal peptides (SPs) pONA, 37f proteolysis of authentic, 14–18 RseP, 11–18 in vitro cleavage assay, 43–44 Site-2 protease (S2P), 128, 208–209, 296–297 expression, 130–131 glimpse, 129–130 metalloproteases, 230 M. jannaschii, 131, 146, 146f purification, 133–135 Size-exclusion chromatography (SEC), 258, 272–274, 275f, 414–415 Solid phase, antigen presentation on, 66–67 Soluble prenyltransferases, 337–341, 338–339f Sonication, 264–265, 324 SPP. See Signal peptide peptidase (SPP) SPP-like family proteases (SPPL), 130

473

Subject Index

SPR. See Surface plasmon resonance (SPR) Staphylococcus aureus, SpsB structure, 38 Staphylococcus epidermidis arylomycin-sensitive, 39 SP1 substrates, 45 Sterol regulatory element-binding protein (SREBP) pathway, 128–130 Streptavidin-Sepharose beads, 179 Substituted-cysteine accessibility method (SCAM), 189–191, 190f, 192f cross-linking experiments, 200–201 using intact cell, 198–199 using microsome, 199–200 Substrates amino-terminal fluorescent label on, 231–233 fluorophore-labeled, 239 inducible proteoliposome assay, 305–306 and intramembrane proteases, 240–242, 244–245, 282–284 orientation in proteoliposomes, 244–245 psTatA, expression and purification of, 401–402 purification, 214–216 specificity of rhomboid protease, 282–285, 283f transmembrane domain, 210–214 Substrate specificity profiling, 38, 46–53, 48f, 53t Surface plasmon resonance (SPR), 81, 85

T Thiol-alkylating reagents, RseP, 18–27 Tobacco etch virus (TEV) protease, 272–274 Transcription activator-like effector nucleases (TALENs)-mediated gene knockout, 370–378, 371–372f Transmembrane domain (TMD) chimeric substrates, 221–222 cleavage of, 296 Drosophila melanogaster, 211–212 intramembrane proteases, 208–209 protein substrates, 117–120 rhomboid protease, 282–285 substrate, 210–214, 222–223

Trichloroacetic acid (TCA), 357 Tris–Bicine urea SDS-PAGE analysis, 166–172

U UBIAD1, 313–314, 339 UbiA superfamily, 310–311 A. pernix, 330–331 Archaeoglobus fulgidus, 315, 330 biological function, 311–315 cloning, 325–326 crystallization and structure determination, 334–337 detergent selection, 333–334 enzymatic activity, 311–315, 319–325 prenyltransferases, 312–313f protein purification, 331–333 sequence clustering analysis, 315–318, 317f structural studies, 325–341 ubiA–menA– BL21(DE3) strain, 323 Ultracentrifugation, 66, 305 Unilamellar liposomes, 238–239

V Virus-like particles (VLPs), 65–66 Vitamin K cycle enzymes cell-based assay, 361–369, 362f ELISA-based evaluation, 362f, 364–366 FIXgla-PC reporter cell lines, 363–364, 365f, 367f Vitamin K-dependent (VKD), 350, 351f carboxylation, 350–351, 351f clotting factors, 350–351 Vitamin K-dependent coagulation factors deficiency (VKCFD), 350–351, 379, 386 Vitamin K epoxide reductase (VKOR) cDNA, 375 DGKO reporter cells, 375–376, 377f dithiothreitol and, 352–354, 370 function study, 352–354, 354f pharmacogenetics, 370 TALENs-mediated gene knockout, 370–378, 371–372f VKOR-Y139F, 368–369 Vitamin K reductase (VKR), 358–361 VKOR homologs (VKORHs), 368–369

474

W “Walker” strains, 102–104 Warfarin enantiomers, 369f

Subject Index

in VKOR mutants, 378 Western blot analysis, 106, 108f, 114f, 140, 144–145, 404–406, 407f Whole cells, 65–66