Redox-Mediated Signal Transduction: Methods and Protocols [2nd ed.] 978-1-4939-9461-8;978-1-4939-9463-2

Table of contents :
Front Matter ....Pages i-x
The Role of Redox in Signal Transduction (John T. Hancock)....Pages 1-11
Methods for the Addition of Redox Compounds (John T. Hancock)....Pages 13-25
Investigating ROS, RNS, and H2S-Sensitive Signaling Proteins (Eleanor Williams, Matthew Whiteman, Mark E. Wood, Ian D. Wilson, Michael R. Ladomery, Joel Allainguillaume et al.)....Pages 27-42
Measurement of 4-Hydroxynonenal (4-HNE) Protein Adducts by ELISA (Kosha Mehta, Vinood B. Patel)....Pages 43-52
Using Flow Cytometry to Detect and Measure Intracellular Thiol Redox Status in Viable T Cells from Heterogeneous Populations (Alex J. Wadley, Rhys G. Morgan, Richard L. Darley, Paul S. Hole, Steven J. Coles)....Pages 53-70
Detection of S-Nitrosation and S-Glutathionylation of the Human Branched-Chain Aminotransferase Proteins (Thomas E. Forshaw, Myra E. Conway)....Pages 71-84
Imaging of Intracellular Hydrogen Peroxide Production with HyPer upon Stimulation of HeLa Cells with EGF (Kseniya N. Markvicheva, Ekaterina A. Bogdanova, Dmitry B. Staroverov, Sergei Lukyanov, Vsevolod V. Belousov)....Pages 85-91
Applications of Electron Paramagnetic Resonance (EPR) Spectroscopy in the Study of Oxidative Stress in Biological Systems (Simon K. Jackson)....Pages 93-102
Extracellular and Intracellular NO Detection in Plants by Diaminofluoresceins (Neidiquele Maria Silveira, Eduardo Caruso Machado, Rafael Vasconcelos Ribeiro)....Pages 103-108
Working with Hypoxia (Elizabeth Bowler, Michael R. Ladomery)....Pages 109-133
Predicting the Effects of Low Dose-Rate Ionizing Radiation on Redox Potential in Plant Cells (Nicol Caplin, Neil Willey)....Pages 135-142
Thioredoxin-1 PEGylation as an In Vitro Method for Drug Target Identification (Jolanta Skalska)....Pages 143-149
Redox-Regulated, Targeted Affinity Isolation of NADH-Dependent Protein Interactions with the Branched Chain Aminotransferase Proteins (Maya E. L. Hindy, Myra E. Conway)....Pages 151-163
Analysis of Redox Relationships in the Plant Cell Cycle: Determination of Ascorbate, Glutathione, and Poly(ADPribose)polymerase (PARP) in Plant Cell Cultures (Christine H. Foyer, Till K. Pellny, Vittoria Locato, Jonathon Hull, Laura De Gara)....Pages 165-181
Equations to Support Redox Experimentation (John T. Hancock, Matthew Whiteman)....Pages 183-195
Back Matter ....Pages 197-199

Citation preview

Methods in Molecular Biology 1990

John T. Hancock Myra E. Conway Editors

RedoxMediated Signal Transduction Methods and Protocols Second Edition

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

For further volumes: http://www.springer.com/series/7651

Redox-Mediated Signal Transduction Methods and Protocols Second Edition

Edited by

John T. Hancock and Myra E. Conway Department of Applied Sciences, University of the West of England, Bristol, UK

Editors John T. Hancock Department of Applied Sciences University of the West of England Bristol, UK

Myra E. Conway Department of Applied Sciences University of the West of England Bristol, UK

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-9461-8 ISBN 978-1-4939-9463-2 (eBook) https://doi.org/10.1007/978-1-4939-9463-2 © Springer Science+Business Media, LLC, part of Springer Nature 2019 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Cover Caption: Knockdown of the redox-active BCAT protein results in increased nuclear localization of MAPK in response to insulin. Image courtesy of: Mai Shafei, Arwa Flembon and David Corry. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 233 Spring Street, New York, NY 10013, U.S.A.

Preface Cellular redox reactions are fundamentally important to bioenergetics, signalling, and overall homeostasis, yet their assessment, particularly in cells, can be challenging. Over the last decade, there has been a drive to develop more sophisticated, sensitive probes to better characterize redox modifications. This book is excellent for those who are not just new to the redox field but also for redox experts that aim to extend their research into new areas or new systems. The importance of developing new methods to measure redox biology is highlighted in the opening chapter, where its role in signal transduction is defined. This chapter covers the main redox components in animal and plant cells with a focus on the role of reactive oxygen species (ROS), reactive nitrogen species (RNS), and hydrogen sulfide in cell signalling. This is coupled with the all-important reducing systems: thioredoxin and glutaredoxin together with the peroxiredoxins, which operate in a fine-tuned response to main homeostasis. The scope of the book is purposely broad to cover a wide application of redox methods, protocols, and applications. The book first introduces the importance of working with redox compounds, in particular the delivery of these reagents. This chapter is an excellent starting point for those new in the field and gives valuable insight into the technical limitations of each redox compound. The advantages and disadvantages of each method are discussed with notes to guide the user on the toxicity of gases such as H2S. Each chapter builds on these compounds where we gain insight into the measurement of these modifications using flow cytometry, ELISA assays, and Western blot analysis (using antibodies specific to 4-hydroxynonenal, S-nitrosation, and S-glutathionylation). These chapters lead nicely to recently developed probes such as the genetically encoded fluorescent probe HyPer, which has been used extensively by experts in the field. The book moves from specific protein modifications to measurement of oxidative stress in mitochondria and biological systems. Here, the importance of electron paramagnetic resonance spectroscopy is highlighted with a step-by-step explanation of experimental design. The detection of NO in plants is also discussed along with the challenges faced when working under hypoxic conditions. We finish the book with several chapters highlighting the broader application of these techniques, which spans from assessment of low-dose-rate ionizing radiation on redox potential to drug target identification, plant cell cycle relationships, and targeting metabolic proteomic partners, where redox active thiols are important to their function. Together these book chapters offer an exciting overview of the scope of cellular redox analysis and the importance of not being limited by technology—new methods and applications are always emerging. We would like to take this opportunity to sincerely thank all the authors for their contribution to this book. These authors are among leaders in their field that have contributed significantly to the assessment of redox cellular biology interactions. Their vision and expertise have contributed to significant advances in our understanding of cell signalling and no doubt with more to come. We are at the beginning of a very exciting chapter of redox

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biology. With the continued commitment to developing new and novel measurements of redox biology, we will be able to fully appreciate the extent and importance of these signalling systems in regulating cellular function, in particular with respect to disease. Finally, we would like to thank the series editor, John Walker, for asking us to edit this book. Bristol, UK

John T. Hancock Myra E. Conway

Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1 The Role of Redox in Signal Transduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 John T. Hancock 2 Methods for the Addition of Redox Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 John T. Hancock 3 Investigating ROS, RNS, and H2S-Sensitive Signaling Proteins . . . . . . . . . . . . . . . 27 Eleanor Williams, Matthew Whiteman, Mark E. Wood, Ian D. Wilson, Michael R. Ladomery, Joel Allainguillaume, Tihana Teklic, Miro Lisjak, and John T. Hancock 4 Measurement of 4-Hydroxynonenal (4-HNE) Protein Adducts by ELISA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43 Kosha Mehta and Vinood B. Patel 5 Using Flow Cytometry to Detect and Measure Intracellular Thiol Redox Status in Viable T Cells from Heterogeneous Populations. . . . . . . . 53 Alex J. Wadley, Rhys G. Morgan, Richard L. Darley, Paul S. Hole, and Steven J. Coles 6 Detection of S-Nitrosation and S-Glutathionylation of the Human Branched-Chain Aminotransferase Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 71 Thomas E. Forshaw and Myra E. Conway 7 Imaging of Intracellular Hydrogen Peroxide Production with HyPer upon Stimulation of HeLa Cells with EGF . . . . . . . . . . . . . . . . . . . . . . 85 Kseniya N. Markvicheva, Ekaterina A. Bogdanova, Dmitry B. Staroverov, Sergei Lukyanov, and Vsevolod V. Belousov 8 Applications of Electron Paramagnetic Resonance (EPR) Spectroscopy in the Study of Oxidative Stress in Biological Systems . . . . . . . . . . . . . . . . . . . . . . . 93 Simon K. Jackson 9 Extracellular and Intracellular NO Detection in Plants by Diaminofluoresceins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 Neidiquele Maria Silveira, Eduardo Caruso Machado, and Rafael Vasconcelos Ribeiro 10 Working with Hypoxia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 Elizabeth Bowler and Michael R. Ladomery 11 Predicting the Effects of Low Dose-Rate Ionizing Radiation on Redox Potential in Plant Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 Nicol Caplin and Neil Willey

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Thioredoxin-1 PEGylation as an In Vitro Method for Drug Target Identification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jolanta Skalska Redox-Regulated, Targeted Affinity Isolation of NADH-Dependent Protein Interactions with the Branched Chain Aminotransferase Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Maya E. L. Hindy and Myra E. Conway Analysis of Redox Relationships in the Plant Cell Cycle: Determination of Ascorbate, Glutathione, and Poly(ADPribose)polymerase (PARP) in Plant Cell Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Christine H. Foyer, Till K. Pellny, Vittoria Locato, Jonathon Hull, and Laura De Gara Equations to Support Redox Experimentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . John T. Hancock and Matthew Whiteman

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors JOEL ALLAINGUILLAUME  Faculty of Health and Applied Sciences, University of the West of England, Bristol, UK VSEVOLOD V. BELOUSOV  Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, Moscow, Russian Federation; Pirogov Russian National Research Medical University, Moscow, Russian Federation EKATERINA A. BOGDANOVA  Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, Moscow, Russian Federation ELIZABETH BOWLER  College of Medicine and Health, University of Exeter Medical School, Exeter, UK NICOL CAPLIN  Faculty of Health and Applied Sciences, University of the West of England, Bristol, UK STEVEN J. COLES  School of Science and the Environment, University of Worcester, Worcester, UK MYRA E. CONWAY  Department of Applied Sciences, University of the West of England, Bristol, UK RICHARD L. DARLEY  Institute of Cancer and Genetics, Cardiff University School of Medicine, Cardiff, UK LAURA DE GARA  Faculty of Biological Sciences, University of Leeds, Leeds, UK THOMAS E. FORSHAW  Faculty of Health and Applied Sciences, University of the West of England, Bristol, UK CHRISTINE H. FOYER  Faculty of Biological Sciences, University of Leeds, Leeds, UK JOHN T. HANCOCK  Department of Applied Sciences, University of the West of England, Bristol, UK MAYA E. L. HINDY  Faculty of Health and Applied Sciences, University of the West of England, Bristol, UK PAUL S. HOLE  Institute of Cancer and Genetics, Cardiff University School of Medicine, Cardiff, UK JONATHON HULL  Faculty of Biological Sciences, University of Leeds, Leeds, UK; Faculty Health and Applied Sciences, University of the West of England, Bristol, UK SIMON K. JACKSON  Faculty of Medicine and Dentistry, Institute of Translational and Stratified Medicine, School of Biomedical Sciences, University of Plymouth, Plymouth, UK MICHAEL R. LADOMERY  Faculty of Health and Applied Sciences, University of the West of England, Bristol, UK MIRO LISJAK  Faculty of Agrobiotechnical Sciences, Josip Juraj Strossmayer University of Osijek, Osijek, Croatia VITTORIA LOCATO  Faculty of Biological Sciences, University of Leeds, Leeds, UK SERGEI LUKYANOV  Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, Moscow, Russian Federation; Pirogov Russian National Research Medical University, Moscow, Russian Federation EDUARDO CARUSO MACHADO  Laboratory of Plant Physiology “Coaracy M. Franco”, Center R&D in Ecophysiology and Biophysics, Agronomic Institute (IAC), Campinas, Brazil KSENIYA N. MARKVICHEVA  Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, Moscow, Russian Federation

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KOSHA MEHTA  School of Applied Sciences, London South Bank University, London, UK RHYS G. MORGAN  School of Life Sciences, University of Sussex, Brighton, UK VINOOD B. PATEL  School of Life Sciences, University of Westminster, Westminster, UK TILL K. PELLNY  Faculty of Biological Sciences, University of Leeds, Leeds, UK RAFAEL VASCONCELOS RIBEIRO  Laboratory of Crop Physiology, Department of Plant Biology, Institute of Biology, University of Campinas (UNICAMP), Campinas, Brazil NEIDIQUELE MARIA SILVEIRA  Laboratory of Plant Physiology “Coaracy M. Franco”, Center R&D in Ecophysiology and Biophysics, Agronomic Institute (IAC), Campinas, Brazil JOLANTA SKALSKA  College of Liberal Arts and Sciences, Alfred University, Alfred, NY, USA; Captor Therapeutics SA, Dunska, Wroclaw, Poland DMITRY B. STAROVEROV  Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, Moscow, Russian Federation TIHANA TEKLIC  Faculty of Agrobiotechnical Sciences, Josip Juraj Strossmayer University of Osijek, Osijek, Croatia ALEX J. WADLEY  School of Sport, Exercise and Health Sciences, Loughborough University, Loughborough, UK MATTHEW WHITEMAN  Medical School Building, University of Exeter, Exeter, UK NEIL WILLEY  Faculty of Health and Applied Sciences, University of the West of England, Bristol, UK ELEANOR WILLIAMS  Faculty of Health and Applied Sciences, University of the West of England, Bristol, UK; Horizon Discovery Ltd., Cambridge, UK IAN D. WILSON  Faculty of Health and Applied Sciences, University of the West of England, Bristol, UK MARK E. WOOD  Geoffrey Pope Building, University of Exeter, Exeter, UK

Chapter 1 The Role of Redox in Signal Transduction John T. Hancock Abstract It is the functioning of efficient cell signaling which is vital for the survival of cells, whether it is a simple prokaryote or a complex eukaryote, including both animals and plants. Over many years various components have been identified and recognized as crucial for the transduction of signals in cells, including small organic molecules and ions. Many of the mechanisms allow for a relatively rapid switching of signals, on or off, with common examples being the G proteins and protein phosphorylation. However, it has become apparent that other amino acid modifications are also vitally important. This includes reactions with nitric oxide, for example S-nitrosation (S-nitrosylation), and, of particular relevance here, oxidation of cysteine residues. Such oxidation will be dependent on the redox status of the intracellular environment in which that protein resides, and this will in turn be dictated by the presence of pro-oxidants and antioxidants, either produced by the cell itself or from the cell’s environment. Here, the chemistry of redox modification of amino acids is introduced, and a general overview of the role of redox in mediating signal transduction is given. Key words Cysteine modification, Hydrogen peroxide, Hydrogen sulfide, Nitric oxide, Redox, Snitrosation, Signal transduction, Thiol modifications

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Introduction Redox chemistry often continues to be intimidating for those that are not passionate about the topic. Students, and indeed researchers, often shy away from lectures or seminars on redox, or quite commonly they simply cannot see the relevance of these principles and formulae for what they are hoping to become in the future. However, when considered at the basic level, redox simply means that there is reduction and oxidation of compounds taking place, that is, there is simply the movement of electrons. Many of the reactions that allow an organism to survive are based on redox. Mitochondria make ATP, our ubiquitous source of useful energy, by harnessing the power of redox. Chloroplasts are driven by redox. Even the mechanisms by which we ward off disease use redox chemistry.

John T. Hancock and Myra E. Conway (eds.), Redox-Mediated Signal Transduction: Methods and Protocols, Methods in Molecular Biology, vol. 1990, https://doi.org/10.1007/978-1-4939-9463-2_1, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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Historically, the world of redox has resided in those laboratories around the world which are interested in energy. How can we take the sun’s energy and make an organism thrive and grow, and how can that healthy lunch be transformed into useful energy to drive our muscles? However, over the years redox has been seen to be more than this, and it was found that redox was being used to rid bodies of xenobiotic materials, via such systems as the P450 complex. Pathogens could be warded off by redox enzymes such as the NADPH oxidase complex, while the blood flow of animals or the transpiration steam of plants can be modulated by the presence of nitric oxide, produced using redox enzymes. Nitric oxide came to the fore in the mid-1980s, and this saw a new era beginning for the redox world. It was found that a molecule previously referred to as endothelial derived relaxing factor (EDRF) was in fact a gaseous free radical molecule, and one that would therefore partake in redox chemistry. That molecule was nitric oxide (NO) and when it was realized that NO was taking part in cell signaling, it opened the door to ask the question about the involvement of redox in general, and other compounds which could impinge on redox chemistry. Many other research groups, who had an interest in the roles of reactive oxygen species (ROS, sometimes referred to as active oxygen species: AOS) started to look to see if manipulation of ROS levels in and around cells had effects on the signaling that ensued, and many discovered that ROS could be considered as key signaling components in many pathways [1–5] (see Note 1).

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Cell Signaling Cell signaling is one of the most important aspects of modern biological sciences. It is clear that for a cell to survive it has to be able to respond to its environment, and change its activity accordingly. This would be just as applicable to a single-celled organism as it would for a cell embedded in a tissue of a mammal. Cells need to respond to environmental cues, whether they are in the form of chemicals in the air or water, or whether they are compounds such as hormones and cytokines generated within the organism itself. Many mechanisms exist to perceive the presence of extracellular factors, for example receptors would be able to detect the arrival of hormones or cytokines, or a protein may recognize a change in environmental conditions such as temperature. Yet further mechanisms are in place to transmit a message deep into the cell once a signal has arrived and initiated a response. The final resultant changes in activity or function may be in a range of places in the cell, including modulation of metabolic activity in the cytoplasm, or the upregulation or downregulation of gene expression in the nucleus (for example see ref. 6).

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No matter what initiates the response, or in fact where the response is finally seen, one of the key mechanisms used by cell signaling pathways is the rapid and reversible turning on or off of protein function. Some cell signaling components have indeed been referred to as molecular switches, such as the G proteins. G proteins can be grouped into two families, heterotrimeric or monomeric, but regardless of this, they can exist in an inactive state, but can be rapidly activated, a process which involves the loss of GDP and the gain of GTP. Intrinsic GTPase activity can then rapidly convert the GTP to GDP, rendering the protein once again inactive. Therefore in the GTP bound state they are able to transmit on a message, but in the GDP bound state will have a different conformation and the ability to interact and pass on a signal will be different. Obviously, this cycle has to be tightly controlled, and many other proteins, including receptors and other interacting partners, are often involved. A more common mechanism for controlling protein activity is the addition of a phosphate group, a process called phosphorylation, which is catalyzed by kinases. Addition of a phosphate group may alter the ability of the protein to have further protein–protein interactions, or may change the conformation of the protein and so alter its catalytic activity. The process can be reversed by removal of the phosphate group, the process of dephosphorylation which is catalyzed by phosphatases. Therefore, it can be seen that evolution has put in place a whole series of ways in which protein function can be quickly altered, with that alteration being able to be undone. The question whether redox chemistry, and the presence of redox compounds, is possibly involved in such signaling pathways therefore arises,

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Redox Mediated Signal Transduction The main compounds which are thought to be involved in redoxmediated signal transduction are quite a large group of chemicals. They include those which are referred to as reactive nitrogen species (RNS) such as nitric oxide and peroxynitrite. A second group are the compounds which come under the umbrella of ROS, and this includes hydrogen peroxide (H2O2), hydroxyl radicals and superoxide anions (O2·–). However, such signaling will also be under the control of other compounds such as glutathione and hydrogen sulfide, and these should be included in discussions too [7]. Indeed, as will be discussed below, both glutathione and hydrogen sulfide (H2S) will be able to compete with RNS and ROS in their interactions with downstream signaling components. It would be foolhardy to consider any of these compounds in isolation as there will be a complex interplay between them which will lead to the ultimate signaling response [8].

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3.1 Redox Molecules as Signals

If NO and ROS, or indeed H2S or glutathione, are to act as signals they need to have certain characteristics [9]. If general guidelines were to be drawn up, then such compounds would need to: – be produced where they are needed; – be produced rapidly and only when needed; – be perceived, and their presence acted upon; – be removed rapidly, so that the signal is not sustained. For both NO and ROS, there appears to be no single source of their production that can be pointed at and deemed to be part of a signaling pathway, and therefore several potential enzymes could be involved. In mammals NO· is primarily produced by a family of nitric oxide synthase (NOS) enzymes, two of which are constitutively present (eNOS and nNOS), and one of which is inducible (iNOS). They are all flavin- and heme-containing enzymes, using Larginine as a substrate, which is converted to citrulline and NO by a short redox pathway. In fact the enzymes have a high degree of similarity to the P450 system, but in NO synthase the reductase domain and the oxidase domain are part of the same polypeptide. However, NO can also be produced by other redox enzymes, such as the molybdenum based xanthine oxidoreductase (otherwise known as xanthine oxidase: XO). In plants, it has been found that another molybdenum-based enzyme produces NO from nitrite, that is, nitrate reductase (NR). Therefore, to implicate any one source of NO to a particular pathway may not be as straightforward as first anticipated (For a review on NO signaling see ref. 10 for example). ROS on the other hand also have a range of sources, and in fact, ROS can arise from many redox pathways. For example, both mitochondria and chloroplasts are known to produce ROS. As well as producing NO, the enzyme xanthine oxidoreductase was first studied as it was known to produce ROS, and many organisms have enzymes that are related to the NAPDH oxidase system. NADPH oxidase was first studied in neutrophils in mammalian host defense, but it is now known that many homologues are involved in ROS production for signaling purposes. In a similar manner, enzymes which can generate H2S have been found in both animals and plants, and so for all these compounds cells have the ability to produce them if they are to be used as signals. By a look at the enzymes involved in the production of NO and ROS, it can be seen that many are themselves controlled in a tight manner, often by the intracellular Ca2+ levels or phosphorylation, and therefore the generation of the relevant redox signaling molecule is controlled well—a good aspect if they are to be considered as part of signal transduction pathways. Furthermore, often the substrate for the reactions are reasonably common, or at least not scarce, materials, such as oxygen, NADPH, nitrite or arginine.

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This means that the production of NO or ROS can usually be rapid and controlled, fulfilling requirements of signaling molecules outlined above. Signaling molecules also need to be removed when no longer needed, and here the very nature of the compounds helps. NO and ROS are inherently unstable. NO is rapidly converted to nitrites and nitrates, while superoxide will dismute to H2O2 especially if the pH is relatively low. Furthermore, cells are replete with antioxidants and antioxidant mechanisms. This includes enzymes such as catalase or superoxide dismutase (SOD) but also includes other compounds such as alpha-tocopherol, glutathione, ascorbate, along with concomitant enzyme systems to recycle such compounds. Therefore, any ROS produced are very likely to be removed very quickly and in fact the rapidly removal of ROS and NO has been used an argument against their involvement in signaling, as it is sometimes unclear how the ROS or NO can reach the proteins that they are potentially controlling. Again, in a similar manner, enzymes have been found that are able to remove H2S (see Note 2). 3.2 ROS and NO Perception

For ROS and NO to be true signals, they need to be perceived by cells, either when they are arriving from the outside, or when they are produced intracellularly. If they are not perceived in some way there will be no response, and no further signaling will take place. It is hard to conceive that there are receptors for NO and ROS that function as “classical” receptors. H2O2 is too small to partake in a ligand-receptor binding event for example. Therefore, it is more likely that ROS and NO use the nature of their chemistry to have their presence felt. In fact, ROS and NO (and H2S for that matter) can react together to alter the signaling potential of each other, which may be one way in which they have a major effect. The most well recognized mechanism for the signaling action of NO is through its reaction with the enzyme guanylyl cyclase (otherwise referred to as guanylate cyclase), promoting the production of cyclic guanosine monophosphate (cGMP or guanosine 30 ,50 -cyclic monophosphate) as depicted in Fig. 1. This can then lead to downstream effects which may involve phosphorylation through the actions of kinases. The exact target on the cyclase for NO is the enzyme’s heme group, and NO can also have an effect on other heme-containing enzymes too, and it has been suggested that some heme-containing enzymes may be involved in modulating NO accumulation in cells [11], such as the hemoglobins. However, both ROS and NO can chemically modify proteins, and the mechanisms involved and the ramifications of such modifications are attracting great interest at the moment.

3.2.1 Covalent Modifications Involving ROS, NO and H2S

One of the main chemical targets for ROS and NO is the thiol group of the amino acid cysteine which may reside within the primary sequence of a protein. As depicted in Fig. 2, there are a

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Fig. 1 NO often has its effects through the activation of guanylyl cyclase

Fig. 2 Cysteine thiol groups can partake in a variety of chemistries, perhaps in competition with each other. Some, but not all, such chemistries are shown here

variety of modifications that are possible on such thiols. Simple oxidation of the thiol in the presence of another not-too-distant cysteine (in three dimensions) may lead to the formation of a disulfide bridge, a simple –S–S– structure (not shown in Fig. 2). The formation of such a bond would potentially both alter the conformation of the protein and stabilize it, allowing for the possible alteration of its activity or function. However, oxidation of a single thiol is also possible, and this may lead to the formation of a sulfenic acid group. In some cases, for example in tyrosine phosphatase, this has been found to rearrange to form the sulfenyl amide group [12, 13]. Further oxidation of the sulfenic acid is also possible and can lead to the formation of a sulfinic acid or further to that of a sulfonic acid (Fig. 2). The more oxidized this thiol becomes, the less likely that it can be re-reduced, and so formation of a

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sulfonic acid is thought to be irreversible, not good for cell signaling, where reversibility is often desired (unless the protein is destined for destruction). On the other hand, reaction of NO with the thiol can lead to the formation of –SNO, a process known as S-nitrosation (also known as S-nitrosylation). The identification of such modifications is the subject of a chapter later in this book (also see ref. 14). However, like oxidation, S-nitrosation can lead to the alteration of activity or function of a protein, Further thiol modification can also take place. Recent interest in the possible signaling role of H2S has shown that thiols can be modified by this gas too, in a process which has been dubbed Ssulfhydration, or more accurately S-persulfication [15]. Cells contain a high concentration of glutathione and this too may react with thiols in the process of S-glutathionylation. Again, all such thiol modification may alter the functionality of the protein, and such chemistry is not rare but used as a potential way of controlling a wide range of proteins [15, 16], and further work in this area will no doubt increase the interest in this biochemistry. Therefore it can be seen that modification of proteins can be the result of exposure to a range of redox compounds such as ROS and NO. Such modifications would potentially be associated with conformational changes in the protein, with the resultant change of activity, either increasing it or decreasing it. This is analogous to the mechanism of phosphorylation, for which there is no doubt about its importance in cell signaling. Of course, if NO and ROS, and even antioxidants such as glutathione, are all vying to react with thiol groups, there will no doubt be a competition between them. It may be that it is the exact concentrations of ROS and NO, and glutathione, that dictate the exact modification that finally takes place. Each type of modification, either S-nitrosation, thiol oxidation, S-persulfication, or Sglutathionylation will potentially have a different effect on the activity or functioning of the protein, and so the activity of NOS, ROS-producing oxidases, and other mechanisms for generating alterations in redox need to be considered in a holistic way. Other amino acid modifications are possible too. NO will react with tyrosine residues in proteins in the process of tyrosine nitration. Methionine modifications in the presence of ROS will lead to oxidation. Of course such alterations of amino acids in proteins will be potentially as important as any other if they can lead to changes in conformation and function. 3.2.2 Proteins Involved in Perception of ROS and NO

One of the challenges in redox signal transduction research has been to identify the proteins involved [17]. Some proteins known to be involved in signal transduction have been found to be inhibited by H2O2, for example tyrosine phosphatase [18], but many other proteins are undoubtedly involved in redox signaling too.

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Using a microarray approach in a study of cells after a treatment of oxidative stress, the ethylene receptor ETR1 was found to be important in H2O2 signalling in Arabidopsis thaliana. Using mutants it was found that a cysteine residue was important for this functioning (Cys65), suggesting that thiol modification is involved, although this has yet to be confirmed ([19], and later in this book). In an alternate approach, thiol tagging with fluorescent markers in the presence and absence of ROS revealed several proteins that could potentially be involved in H2O2 signaling, including glyceraldehyde 3-phosphate dehydrogenase (GAPDH: [20]), a protein more usually associated with metabolism, in particular the glycolytic pathway. Interestingly, once modified this protein translocates to the nucleus where it partakes in the alteration of gene expression, a role far different than its enzymatic role in the cytoplasm. Gene expression alterations are not uncommon in redoxmediated signaling. Transcription factors have also been identified that are controlled in a redox-mediated manner. In Escherichia coli, for example, the transcription factor OxyR is activated by H2O2, leading to disulfide bond formation between cysteine 199 and cysteine 208 [21]. However, it is not only ROS for which proteins which are altered have been identified. Using methods such as those proposed by Snyder and Jaffrey [22] proteins which undergo S-nitrosation have been identified, and these are wide ranging, including many which are involved in cell signaling events. Similarly, proteins which have undergone S-persulfidation [15] have also been started to be identified, and such methodologies are further discussed later in this book. Therefore it can be seen that there are a wide variety of proteins that might take part in the perception of redox and redox active molecules, including ones that are already associated with intracellular signal transduction pathways, ones that are involved in other activities such as metabolism, and others that are in direct control of gene expression. The challenge remains to untangle how this all functions and is coordinated, especially considering that these redox active molecules are often produced in response to the same cues. 3.2.3 Systems Involved in Reversal of Redox Modifications

Once a cysteine thiol has been oxidized, it needs to be re-reduced if the signaling is to be ablated. If the disulfide bond has been created by ROS reduction, thioredoxins (TRXs), and glutaredoxins may act as protein disulfide reductases [23, 24]. TRXs and glutaredoxins may also reoxidize –SOH groups [25, 26]. If ROS oxidation has caused the formation of the sulfinic acid group, this can be reduced back to the sulfenic acid group by sulfiredoxins. These ATP-dependent enzymes were first identified in yeast [27]. The sulfenic acid group created can then be further reduced by TRX or

The Role of Redox in Signal Transduction

9

glutaredoxins resulting in the regeneration of the thiol, –SH, and so allowing a further round of signaling. In a similar manner, S-NO groups can be reduced back to the thiol group by enzymes such as SNO reductases, therefore allowing such proteins to be reprogrammed for a further round of signaling [28, 29].

4

Conclusion There are many compounds, including ROS, RNS, and sulfurbased compounds, from a variety of sources, that can influence the redox status of cells, and furthermore these can potentially interact with each other or modify a range of proteins via redox chemistry. Many of these changes to such proteins will result in alterations of their activities and functions, and therefore a greater understanding of how redox is controlled and how changes in redox influence metabolic pathways or the expression of genes is required to fully elucidate how redox chemistry fits into the overall scheme of cell signaling. The chapters that follow in this book will discuss the influence of aspects of redox chemistry in more depth, and discuss some of the key methodologies that may be employed to study this growing area of research. For a more full text on cell signaling in general and the components involved see refs. 8, 30.

5

Notes 1. A note of caution is added here. The notation for nitric oxide of NO is used in this text. However, it should be noted that the radical form of NO should be written as NO·. Furthermore, with gain or loss of electrons nitric oxide can also be NO and NO+. It cannot be assumed that NO donors release NO·, some release NO+ for example, so when NO donors are used the exact chemistry needs to be checked and understood before data is interpreted. 2. It is the efficient removal of ROS which underpins the advice to eat fruit and vegetables. In the UK the advice is five-a-day, but this is much higher in some parts of the world, especially Scandinavia and Japan.

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References 1. Hancock JT (1997) Superoxide, hydrogen peroxide and nitric oxide as signalling molecules: their production and role in disease. Br J Biomed Sci 54:38–46 2. Dro¨ge W (2002) Free Radicals in physiological control of cell function. Physiol Rev 82:47–95 3. Neill SJ, Desikan R, Hancock JT (2002) Hydrogen peroxide signalling. Curr Opin Plant Biol 5:388–395 4. Colavitti R, Finkel T (2005) Reactive oxygen species as mediators of cellular senescence. IUBMB Life 57:277–281 5. Hancock JT (2009) The role of redox mechanisms in cell signaling. Mol Biotechnol 43:162–166 6. Vanderauwera S, Zimmermann P, Rombauts S, Vandenbeele S, Langebartels C, Gruissem W, Inze´ D, Van Breusegem F (2005) Genomewide analysis of hydrogen peroxide–regulated gene expression in Arabidopsis reveals a high light-induced transcriptional cluster involved in anthocyanin biosynthesis. Plant Physiol 139:806–821 7. Ullrich V, Kissner R (2006) Redox signaling: bioinorganic chemistry at its best. J Inorg Biochem 100:2079–2086 8. Hancock JT, Whiteman M (2014) Hydrogen sulfide and cell signaling: team player or referee? Plant Physiol Biochem 78:37–42 9. Hancock JT (2016) Cell Signalling, 4th edn. Oxford University Press, Oxford 10. Neill SJ, Desikan R, Hancock JT (2003) Nitric oxide signalling in plants. New Phytol 159:11–35 11. Perazzolli M, Romero-Puertas MC, Delledonne M (2006) Modulation of nitric oxide bioactivity by plant haemoglobins. J Exp Bot 57:479–488 12. Salmeen A, Anderson JN, Myers MP, Meng TC, Hinks JA, Tonks NK, Barford D (2003) Redox regulation of protein tyrosine phosphatase 1B involves a novel sulfenyl-amide intermediate. Nature 423:769–773 13. Van Montfort RL, Congreve M, Tisi D, Carr R, Jhoti H (2003) Oxidation state of the active-site cysteine in protein tyrosine phosphatase 1B. Nature 423:773–777 14. Lindermayr C, Saalbach G, Durner J (2005) Proteomic identification of S-nitrosylated proteins in Arabidopsis. Plant Physiol 137:921–930 15. Sen N, Paul BD, Gadalla MM et al (2012) Hydrogen sulfide-linked sulfhydration of

NF-κB mediates its antiapoptotic actions. Mol Cell 45:13–24 16. Dixon DP, Skipsey M, Grundy NM, Edwards R (2005) Stress-induced protein S-glutathionylation in Arabidopsis. Plant Physiol 138:2233–2244 17. Hancock J, Desikan R, Harrison J, Bright J, Hooley R, Neill S (2006) Doing the unexpected: proteins involved in hydrogen peroxide perception. J Exp Bot 57:1711–1718 18. Cho S-H, Lee C-C, Ahn Y, Kim H, Yang K-S, Lee S-R (2004) Redox regulation of PTEN and protein tyrosine phosphatase in H2O2mediated cell signalling. FEBS Lett 560:7–13 19. Desikan R, Hancock JT, Bright J, Harrison J, Weir I, Hooley R, Neill SJ (2005) A novel role for ETR1: hydrogen peroxide signalling in stomatal guard cells. Plant Physiol 137:831–834 20. Hancock JT, Henson D, Nyirenda M, Desikan R, Harrison J, Lewis L, Hughes J, Neill SJ (2005) Proteomic identification of glyceraldehyde 3-phosphate dehydrogenase as an inhibitory target of hydrogen peroxide in Arabidopsis. Plant Physiol Biochem 43:828–835 21. Lee C, Lee SM, Mukhopadhyay P, Kim SJ, Lee SC, Ahn WS, Yu MH, Stroz G, Ryu SE (2004) Redox regulation of OxyR requires specific disulfide bond formation involving a rapid kinetic reaction path. Nat Struct Mol Biol 11:1179–1185 22. Zhang Y, Keszler A, Broniowska KA, Hogg N (2005) Characterization and application of the biotin-switch assay for the identification of Snitrosated proteins. Free Radic Biol Med 38:871–881 23. Schu¨rmann P, Jacquot JP (2000) Plant thioredoxin systems revisited. Annu Rev Plant Physiol Plant Mol Biol 51:371–400 24. Lemaire SD (2004) The glutaredoxin family in oxygenic photosynthetic organisms. Photosynth Res 79:305–318 25. Collin V, Lankemeyer P, Miginiac-Maslow M, Hirasawa M, Knaff DB, Dietz KJ, IssakidisBourguet E (2004) Characterization of plastidial thioredoxins belonging to the new y-type. Plant Physiol 136:4088–4095 26. Rouhier N, Gelhaye E, Sautiere PE, Brun A, Laurent P, Tagu D, Gerard J, De Fay E, Meyer Y, Jacquot JP (2001) Isolation and characterization of a new peroxiredoxin from poplar sieve tubes that uses either glutaredoxin or thioredoxin as a proton donor. Plant Physiol 127:1299–1309

The Role of Redox in Signal Transduction 27. Biteau B, Labarre J, Toledano MB (2003) ATP-dependent reduction of cysteinesulphinic acid by S. cerevisiae sulphiredoxin. Nature 425:980–984 28. Malik SI, Hussain A, Yun BW, Spoel SH, Loake GJ (2011) GSNOR-mediated de-nitrosylation in the plant defence response. Plant Sci 181:540–544

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29. Yu M, Yun BW, Spoel SH, Loake GJ (2012) A sleigh ride through the SNO: regulation of plant immune function by protein S-nitrosylation. Curr Opin Plant Biol 15:424–430 30. Hancock JT (2003) The principles of cell signalling. In: Kumar S, Bentley PJ (eds) On Growth, Form and Computers. Academic, London

Chapter 2 Methods for the Addition of Redox Compounds John T. Hancock Abstract Often in redox biology experiments there is a need to add compounds which impinge on the redox of the cellular environment cell. Such compounds may include reactive oxygen species (ROS), such as hydrogen peroxide (H2O2), reactive nitrogen species such as nitric oxide (NO), hydrogen sulfide (H2S), or even hydrogen gas (H2). It is not always easy or obvious how such compounds should be used. Gases may be supplied and used in the gaseous form, but this is often not convenient. Alternative methods may involve donor molecules that release into solution the relevant compound, but the actual compound released needs to be considered, along with the kinetics of that release and the by-products that might be remain. Therefore, the method of delivery of redox active compounds needs to have careful consideration before more complex experiments are undertaken. This chapter covers some of the more common methods employed and discusses some of the pros and cons of such methods. Key words Hydrogen gas, Hydrogen peroxide, Hydrogen sulfide, Nitric oxide, Reactive nitrogen species, Reactive oxygen species, Superoxide anions.

1

Introduction It is now generally accepted that developmental mechanisms and many stress responses in a variety of organisms involve or are mediated by the generation of reactive compounds which impinge on cell signaling events [1, 2]. Such reactive compounds include reactive oxygen species (ROS) such as the superoxide anion (O2·
) and hydrogen peroxide (H2O2), the reactive nitrogen species such as nitric oxide (NO: although this should be written as the radical form NO·, which is usually the bioactive compound), as well as other compounds such as hydrogen sulfide (H2S) and possibly hydrogen gas (H2). Often, these are generated at the same time in cells and together will orchestrate the appropriate response [3, 4]. ROS can be generated in cells in a variety of ways. In the presence of oxygen, such as in an aerobic organism, electron

John T. Hancock and Myra E. Conway (eds.), Redox-Mediated Signal Transduction: Methods and Protocols, Methods in Molecular Biology, vol. 1990, https://doi.org/10.1007/978-1-4939-9463-2_2, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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leakage from electron transport chains can result in the accumulation of superoxide and H2O2. Alternatively, there are dedicated enzymes which produce ROS, such as the NADPH oxidase enzymes (Nox family of proteins), xanthine oxidoreductase, and peroxidases. Such enzyme activities are usually under tight control so that the generation of ROS is only where and when needed. The generation of superoxide will lead to its dismutation, producing H2O2, and in the presence of metal ions, in the Fenton reaction, this can lead to the generation of the hydroxyl radical (·OH), thought to be the most reactive biomolecule in cells. Further by-products of ROS metabolism can include singlet oxygen (O21), hypochlorous acid (HOCl), or in interaction with NO peroxynitrite (ONOO
). Some of the early work on ROS concentrated on the deleterious effects of ROS, and how such chemistry could be used to protect organisms from pathogen challenge, in both animals and plants [5]. More recently, the positive roles of ROS in cells have been recognized [1, 2]: in particular H2O2 acts as a cell signaling molecule. Downstream events in ROS signaling include the covalent modification of proteins, leading to altered phosphorylation levels in cells and modulated gene expression. Nitric oxide (NO) came to prominence due to its role as endothelial derived relaxing factor (EDRF) [6]. In humans there are three enzymes that generate NO: endothelial nitric oxide synthase (eNOS), neuronal nitric oxide synthase (nNOS), and inducible nitric oxide synthase (iNOS). In plants there is probably no NOS enzyme [7] but NO is produced by nitrate reductase [8]. Downstream NO reacts with iron-based proteins such as hemoglobin but also causes the covalent modification of proteins by a process known as S-nitrosation (often referred to as S-nitrosylation). There are a myriad of responses in both animals and plants, including the control of smooth muscles cells and therefore blood flow, and the closure of leaf stomata. In a similar manner, H2S has been shown to alter blood flow in mammals and is involved in a range of stress responses. There are enzymes that produce it, and it is involved in a range of developmental and stress responses in plants, from seed germination to post-harvest [9], while in lower animals it has been shown to bestow thermotolerance and increase life spans [10]. Lastly, but certainly not insignificantly, the effects of H2 gas have been investigated, and H2 gas has been mooted as a new potential signaling molecule [11]. It has been shown to have effects in animals and plants, and has been suggested to be a future important molecule to be used in therapeutic treatments [12] and in agriculture [13]. It is clear therefore that biological systems, whether they are whole organisms, tissues, cells, or proteins, are able to be modulated by the presence of a range of reactive chemical species.

Addition of Redox Compounds

15

Therefore, experiments are often designed to emulate the environments to which such systems are exposed, that is, in the presence of ROS, NO etc. There is a need to create conditions in which such reactive chemical species can be delivered to the experimental system being mooted. This may be in a variety of media. For example, plants may be exposed to chemicals in the air, or mixed with other gases. Cells in culture may need delivery via the medium in which the cells are growing, while enzymatic assays may require the delivery in assay buffers. To this end, there have been a variety of methods for delivery of ROS, NO, H2S and H2. NO donors are commonly used [14], while some recent work has led to the development of donor compounds for H2S [15, 16]. Some of the more common delivery methods are discussed below, along with some of the considerations which may be needed before the final experimental design is decided upon.

2

Materials

2.1 Reactive Oxygen Species

1. Potassium superoxide (KO2) can be purchased in solid form from commercial companies (e.g., Sigma-Aldrich).

2.1.1 Delivery of ROS in Chemical Form

2. Hydrogen peroxide (H2O2: see Note 1) can be purchased as a solution (e.g., from Sigma-Aldrich or even local pharmacies).

2.1.2 Delivery of Superoxide from Coculture with Neutrophils

1. 0.9% (w/v) NaCl containing EDTA (1 mM) and 5 i.u. heparin/mL. 2. Catering gelatin. 3. 0.2% saline containing EDTA (1 mM) and 5 i.u. heparin/mL. 4. 1.6% saline containing EDTA (1 mM) and 5 i.u. heparin/mL. 5. Ficoll (type F-P). 6. Phorbol 12-myristate 13-acetate (PMA). 7. Suitable resuspension buffer, which will be determined by the final use of the neutrophils. This may be PBS (see Note 2), Krebs-Ringer buffer or tissue culture medium. The alternate method uses the same solutions but recommends the use of Histopaque.

2.1.3 Delivery of ROS Using Enzymes

1. Xanthine oxidase: 1 U/mL needed as final concentration in working buffer/medium. 2. Xanthine: stock solution in working buffer, for example PBS or cell culture medium. Final concentrations up to 1 mM. 3. If testing of ROS production is anticipated then nitroblue tetrazolium (NBT) will also be required.

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John T. Hancock

Nitric Oxide

2.2.1 Delivered as a Gas 2.2.2 Delivered Using Donors

Pressurized gas bottles of NO can be purchased commercially (see Note 3). These can be found in different sizes and can be obtained from BOC Ltd. (see Note 4). 1. Numerous NO donor molecules, such as listed here, can be purchased commercially, for example from Sigma-Aldrich. (a) Sodium nitroprusside (SNP). (b) S-Nitrosyl-N-Acetyl-DL-penicillamine (SNAP). (c) S-Nitrosoglutathione (GSNO).

2.3

Hydrogen Sulfide

2.3.1 Delivered as a Gas

2.3.2 Delivered from Donors

Pressurized gas bottles of H2S can be purchased commercially (see Note 3). These can be found in different sizes and different mixtures with other gases, and can be obtained from BOC Ltd. (see Note 4). 1. Quick release compounds such as sodium sulfide (Na2S) and sodium hydrosulfide (NaSH). 2. Donors such as GYY4137 and AP39 can be purchased from several commercial outlets.

2.4

Hydrogen Gas

2.4.1 Delivered as a Gas 2.4.2 Delivered in Solution

3

Pressurized gas bottles of H2 can be purchased commercially (see Note 3). These can be found in different sizes and can be obtained from BOC Ltd. (see Note 5). “Hydrogen tablets” can be purchased from a variety of sources, such as I Love Hydrogen (Alkaway, UK), Active H2® ULTRA All-Natural Molecular Hydrogen Antioxidant (integratedhealth. com). Alternatively, drinking bottles which purport to generate hydrogen gas can be purchased, for example from naturesenergieshealth.com.

Methods

3.1 Delivery of Reactive Oxygen Species 3.1.1 Delivery of Superoxide Anions

1. Superoxide anions can be delivered directly into solution by dissolving potassium superoxide (KO2) into the appropriate solution (i.e., the buffer or growth media in which biological material is being placed) and then using immediately. A concentrated stock solution from which aliquots can be used should be prepared immediately before use. 2. Alternatively superoxide can be delivered from coculture with cells which generate it. Cells such as neutrophils have a relatively high amount of the enzyme NADPH oxidase which can be activated by the addition of a phorbol ester (phorbol myristate acetate: PMA (see Note 6)). Neutrophils can be prepared

Addition of Redox Compounds

17

from whole blood. For large scale neutrophil isolation the following protocol is effective [17] (see Note 7): (a) Collect whole blood into in 0.9% (w/v) NaCl containing EDTA (1 mM) and 5 i.u. heparin/mL to prevent coagulation. (b) Add catering gelatin to give a final concentration of 1.25% (w/v) and mixed carefully. (c) Leave to stand for about 1 h. (d) Aspirate the top layer which is enriched in leucocytes, leaving the majority of red blood cells behind. (e) Centrifuge the cells (400  g), which will create a cellular pellet. (f) Wash the cells with 0.9% (w/v) NaCl containing EDTA (1 mM). (g) Mix gently, to avoid activation (see Note 7). (h) To remove the remaining red blood cells the cell mixture needs to be exposed to an osmotic shock. To do this, add five volumes of 0.2% saline containing EDTA (1 mM) and 5 i.u. heparin/mL. Mix gently. (i) Leave for 30 s. (j) Add an equal volume of 1.6% saline (with EDTA and heparin). Mix gently. (k) Centrifuge the cells (100  g), which will create a cellular pellet. (l) Repeat the osmotic shock step two more times, unless neutrophils look clean (i.e., not tainted red). (m) Once white blood cell fraction is clean from red cells resuspend in 0.9% NaCl containing 1 mM-EDTA and 5 i.u. of heparin/mL. (n) Overlay on Ficoll. Do this extremely carefully so that the layers are not mixed. It is crucial to minimize mixing of the solutions so position the pipette tip as close to the meniscus of the separation solution as possible and raise the tip as pipetting proceeds. (o) Centrifuged at 400  g for 20 min. (p) Remove the supernatant and keep the pellet. (q) Wash the cells in 0.9% saline (containing EDTA and heparin (5 i.u./mL)) (r) Finally resuspend cells in required buffer (e.g., PBS, or Krebs-Ringer buffer) or tissue culture medium. (s) To activate the neutrophils add 1 μg/mL PMA (see Note 6). Once activated, the NADPH oxidase of the

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neutrophils will generate superoxide anions from molecular oxygen. At pH 7.0, dismutation will lead to the accumulation of H2O2 and downstream ROS by-products. An alternate method has more recently been published [18] which is ideal for smaller volumes of neutrophil isolation. Briefly: (a) An aliquot of whole blood should be carefully layered over an equal volume of separation medium, for example Histopaque. This contains polysucrose and sodium diatrizoate, which has a density of 1.119 g/mL. Overlay the cells carefully as described above. (b) Centrifuge at 500  g for 30–35 min at room temperature. (c) Layers of cells should be visible. Remove cells that are in the layer and most of the solution. Neutrophils and red blood cells are in the pellet. (d) Once again the red blood cells need to be removed. To do this the cells need to be hydrolyzed in hypotonic solution. Resuspend the pellet in 0.2% saline solution for a few seconds (less than 30 s), and then add 1.6% saline to mix briefly. (e) Centrifuge the solution for 250  g for 5–10 min. Remove the red supernatant. This step may need to be repeated to remove the majority of red blood cells. (f) Resuspend final pellet in PBS or cell culture medium. Neutrophils need to be used fresh as they have a limited life-time (1–2 h). 3.1.2 Delivery of Hydrogen Peroxide Delivery as a Solution

Delivery Using Enzymes

Hydrogen peroxide can be delivered as a solution to cells and tissues (see Note 1). Usually a working stock solution is made which is 10–20 more concentrated than required is made and aliquots of this used to ensure a final working concentration is as needed. Often the original concentration as delivered from commercial suppliers is 30% w/w H2O2. One of the easiest ways to deliver ROS to solutions is to use the enzyme xanthine oxidase [19]. To do this: 1. For the cell culture or biological material under study ensure a working concentration of 1 U/mL of xanthine oxidase. This can be added straight to the solution when required. 2. When the generation of ROS is required add a working concentration of up to 1 mM xanthine. A report by Uy et al. [19] shows using the nitroblue tetrazolium (NBT) assay that ROS production was linear between 0 and 1000 μM xanthine.

Addition of Redox Compounds

19

3. For long-term treatments the xanthine will need to be added repetitively over time as the production of ROS will follow Michaelis–Menten kinetics and fall as the substrate concentration goes down. The time frame of this may be a 10–30 min.

3.2.1 Delivery of Nitric Oxide as a Gas

As a gas NO can be delivered straight to a biological material, such as a plant, but it is toxic and care should be taken (see Note 8). Alternatively, NO does dissolve in water [20] so a saturated aqueous solution may be made. Aliquots of this can then be added straight to experiments. To make a saturated solution NO from a pressurized gas cylinder can be bubbled through a solution in a glass container, but this does need to be vented safely (see Note 8).

3.2.2 Delivery Nitric Oxide from Donor Molecules

As NO is not easy, or indeed very safe, to use as a gas many donor molecules have been used in biological experiments over the years [14]. To use such donors:

3.2 Delivery of Nitric Oxide

1. Make an appropriate aqueous buffer for the experiment. 2. Dissolve the NO donor in the buffer at concentrations higher (perhaps tenfold) than required. 3. Pipette aliquots of the stock solution made to the experiments as needed to achieve the final working concentration. NO donors have been classified into groups [14]: nitrovasodilators; S-nitrosothiols; NONOates. Taking representative examples of these, some of the NO donors commonly used are discussed below: 1. Nitrovasodilators: (a) Sodium nitroprusside (SNP) is one of the most commonly used. This is a white solid which releases NO+. Therefore, care needs to be taken in interpreting data here, as it is not the NO· free radical form that is being donated. However, there is another drawback here too, as one of the by-products is cyanide [21]. Cyanide is extremely toxic, but the toxicity illustrates the problem—it is an efficient inhibitor of several important enzyme systems, such as mitochondrial electron transfer so interfering with ATP production, and superoxide dismutase (SOD) so interfering with the antioxidant capacity of cells. Cyanide can also have biological effects in its own right, as well as its toxicity so the data can be very difficult to interpret in this instance [21]. 2. S-Nitrosothiols. These can be adducts of cysteine, N-acetylcysteine, and commonly reduced glutathione (GSH). Concentrations of stock solutions can be easily determined using absorbance at 334 nm (ε ¼ 908 M
1 cm
1, [14]).

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(a) S-Nitrosoglutathione (GSNO). . Although this is a biomolecule which is thought to be instrumental in the transport of NO around organisms, as a pure compound it can be extremely useful for the delivery of NO into solution. One drawback here is that the NO generation by compounds like this is light dependent. In the presence of light, NO release is very greatly enhanced [14]. These compounds also have limited storage times, even at 0  C and in the dark [14]. As discussed further below, the residual by-product needs to be considered here as it is likely to have bioactivity. (b) S-Nitrosyl-N-Acetyl-DL-penicillamine (SNAP). This is another S-nitrosothiols that is commonly used in biological systems, and is in fact a potent vasodilator. It is green in color and soluble in dimethylsulfoxide (see Note 9), which should be used to make stock solutions. 3. NONOates. NONOates, or more correctly named diazeniumdiolates, are NO adducts of various parent compounds such as diethylamine. Concentrations of stock solution can be assayed using the extinction coefficient at 250 nm (ε ¼ 8000 M
1 cm
1). Degradation of, and therefore release of NO from, NONOates is very pH sensitive: a change of approximately 0.5 pH units can alter rates as much as threefold. Long-term storage of these compounds is not recommended. The by-products will be the parent compound, which is an amine and therefore may well be bioactive. To make control solutions overnight decomposition at 37  C in the desired assay buffer is recommended [14]. Lastly, it should be noted that the decomposition rates are very dependent on the parent compound. Half-lives have been quoted as follows: diethylene/NO (DEA/NO), 2–4 min; N-propyl-1-3-propanediamine/NO (PAPA/NO), 15 min; Spermine/NO, 39 min; diethylenetriamine/NO (DETA/ NO), 20 h. 4. Nitro-fatty acids: New NO donors are being suggested on a regular basis and one of the most recent is based in lipid metabolism. It is suggested that cells create nitro-fatty acids, and that these can also be used for NO delivery [22]. These could be good for targets which are relatively hydrophobic locations in the cell. Donor molecules will release the gas into solution but then leave behind a residue by-product. This residue material may be bioactive and therefore this needs to be controlled for. To do this: 1. Make a stock solution of the donor molecule. 2. Leave open to the air for 1–2 h (depending on the kinetics of the gas release of the donor molecule. Overnight is often a good time to ensure donors are fully depleted).

Addition of Redox Compounds

21

3. Use in control experiments in aliquots which are exactly equivalent to the concentrations used for NO treatment experiments. Data from these treatments can be used as control background levels of response or activity. 3.2.3 Use of More Than One Donor and Measurement of NO

Although not covered by this chapter, it is always worth checking the presence of NO when donors are being used. It has been suggested that more than one method is used to confirm the presence of NO, as the specificity of some reactions is still in doubt [23]. The same argument can be said about NO donors and therefore experiments should be repeated with a second independent donor as an ideal.

3.3 Delivery of Hydrogen Sulfide (H2S)

As with other gaseous biomolecules H2S can be delivered as a gas. Samples can be placed in container and gas added straight from a pressurized bottle. However, besides being extremely toxic—it inhibits mitochondrial electron transfer at complex IV—it has an extremely strong odor. Extremely low concentrations in air can be sensed by the human nose, giving a rotten egg smell. Therefore, using H2S as a gas is often avoided, where possible. If safety precautions are being taken appropriately H2S should not be smelt in the laboratory.

3.3.1 Delivery of Hydrogen Sulfide as a Gas

3.3.2 Delivery of H2S from Donors

Several solid donors can be purchased which release H2S into solution. A review of many such donors was written by Zhao et al. [24]. Two of the most commonly used donors are sodium sulfide (Na2S) and sodium hydrosulfide (NaSH). To use such donors: 1. Decide on the appropriate aqueous buffer for the experiments being undertaken (e.g., phosphate buffered saline (PBS)) (see Note 2). 2. Make stock solution of the donor, usually tenfold stronger than the final concentration needed. 3. Add the appropriate volume of donor stock solution to the experiment at the appropriate time to give the final working concentration needed. It should be noted here that H2S is of course a gas and will return to the gaseous phase quickly, as it is not very soluble [25]. Therefore, repeat applications of the solution may be required. Further, once returned to the gaseous phase it can be sensed by the human nose so experiments may need to be carried out in a fume hood. The kinetics of the delivery of H2S by donors such as above does create a problem, as the biological system under study will be exposed to a relatively high concentration of the gas—both NaSH and Na2S release H2S into solution rapidly—for a short period of

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time after which exposure to the gas is limited, as it has gone into the atmosphere. Therefore alternative new donors which release H2S gas in a slower manner, more akin to what may be physiological have been developed. The first of these was a compound called GYY4137 [15] which has been used quite extensively in such studies [26, 27]. Since its development other compounds have been created which are targeted to specific cellular organelles, such as the mitochondria. One such compound is AP39 [16]. 1. To use all such H2S donors, a stock solution needs to be made which is more concentrated than needed and then aliquots added to experiments to get the final working concentrations. As with all donors, there is a by-product remaining at the end after the release of the H2S gas (see discussion above for NO donors). Therefore, suitable controls need to be carried out with depleted donors to ensure biological effects are not from the residue products. 3.3.3 Use of More Than One Donor and Measurement of H2S

As with donors of other redox compounds, it is always good practice to use more than one donor, especially if quick release donors such as NaSH are used. Experiments should be repeated with a second independent donor, such as GYY4137. Also, as discussed briefly with NO, the release of the active compound— here H2S—from the donor can be measured. Again outside the scope of this chapter but methods for the measurement of H2S are used [28].

3.4 Delivery of Hydrogen Gas (H2)

As a gas molecular hydrogen is easy to deliver. Biological systems can be sealed, in a gas container for example, and the atmosphere flooded with gas from a pressurized bottle. For some systems, such as plants, it may be desirable to have a high H2 concentration, although this will of course preclude oxygen from being present. H2 can be purchased as different mixtures, or a mixing valve may be used to regulate the correct H2/air or H2 nitrogen mix. Such an approach was used by Renwick et al. [29] when they studied seeds germination and added into the mix small amounts of oxygen after a 24-h H2 treatment.

3.4.1 Delivery of Molecular Hydrogen as a Gas

3.4.2 Delivery of Hydrogen Gas in Solution

Often it is easier and more desirable to deliver the H2 gas in solution, although it is not very soluble. It has been quoted that saturation in water at 1 atm pressure is 0.78 mmol/L or 1.57 mg/L [30]. Experiments are often carried out using solutions which have been bubbled with hydrogen to create what is referred to as hydrogen-rich water (HRW) and hydrogen-rich saline (HRS), if a saline solution is used to start with (this would normally be isotonic saline which is 0.90% w/v of NaCl).

Addition of Redox Compounds

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Alternatively, there are other methods to create HRW. One is to purchase a hydrogen generating drinking bottle. Here, the bottle is simply filled with water and the generation of hydrogen gas takes place inside. After a period of time, dictated by the manufacturer, the solution is ready to use (it is supposed to be drunk but equally can be used as an experimental solution). Equally easy is the use of hydrogen generating tablets. To do this: 1. Fill a suitable and sealable container with 500 mL of water (double distilled or equivalent should be used for experimental work). 2. Add one hydrogen generating tablet. 3. Seal the container for 10 min. 4. Use the solution immediately the container is unsealed. It should be noted here that as with many gases in solution, H2 will rapidly return to the gaseous phase, that is, the atmosphere and therefore the concentration of H2 in solution will rapidly decline if the solution is not sealed. 1. For disposal of the solution after the experiment it has to be remembered that it is a flammable gas (see Note 5). Solutions can be left to degas in a suitable fume hood, but the solutions are inherently nontoxic and therefore can be disposed of in the normal drainage system if diluted with excess water (see Note 8). Two final points are needed here. Many of the hydrogengenerating tablets are not pure for the chemistry which creates the hydrogen, and they contain other compounds. For example, I Love H2 tablets contain maltose, malic acid, magnesium oxide, magnesium malate, fumaric acid as well as magnesium. Many of these chemicals will have bioactivity in their own right; therefore, controls of hydrogen-depleted solutions (that are left for a period of time so that the hydrogen is no longer in solution but the other factors are) need to be considered. Secondly, as the hydrogen rapidly leaves solution, multiple treatments over time will be needed if the experiments are left for a period of time. It is possible that one treatment leaves long-term effects in some cells, perhaps altering phosphorylation levels, but in some systems regularly repeated treatments may be required.

4

Notes 1. Although hydrogen peroxide can be purchased easily from commercial outlets, including pharmacies, it is often used as a terrorist weapon and therefore needs to be kept in a secure place.

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2. Phosphate buffered saline (PBS). This can be purchased in tablet form (e.g., from Sigma-Aldrich), or created in the laboratory from component materials (1 L PBS contains: 8 g NaCl; 0.2 g KCl; 1.44 g Na2HPO4; 0.24 g KH2PO4; adjusted to pH 7.4 with HCl). 3. Extreme caution should be taken when using pressurized gas bottles. The correct regulators must be fitted and they must be strapped to benches when in use. Risk assessments must also include their movement and transport. 4. Some gases such as nitric oxide and hydrogen sulfide are extremely toxic and must not be breathed in. See for example the BOC safety sheets for: Nitric oxide: https://www.boconline.co.uk/internet.lg.lg.gbr/en/images/ sg-088-nitric-oxide-v1.2410_39637.pdf?v¼3.0. Hydrogen sulfide: https://www.boconline.co.uk/internet.lg.lg.gbr/en/images/ sg_073_hydrogen_sulphide_released410_81719.pdf?v¼4.0. 5. Hydrogen gas is extremely flammable. 6. Phorbol myristate acetate (PMA) is extremely toxic. It acts as a cocarcinogen and should be treated with caution. As a minimum a laboratory coat, gloves, and eye protection should be worn when using PMA. 7. Care needs to be taken preparing neutrophils as they can easily be activated, and then no use for further experiments. Therefore, mixing and aspiration needs to be done carefully, and the correct solutions used. If cells start to aggregate then this is a sign that they have already been activated. 8. Many of these compounds and solutions are toxic or flammable, and always need to be used under appropriate risk assessments. 9. If dimethylsulfoxide (DMSO) is used as a solvent, the equivalent DMSO concentrations need to be used in control experiments, as DMSO has its own bioactivity. References 1. Hancock JT (2009) The role of redox mechanisms in cell signalling. Mol Biotechnol 43:162–166 2. Hancock JT (2017) Harnessing evolutionary toxins for signaling: reactive oxygen species, nitric oxide and hydrogen sulfide in plant cell regulation. Front Plant Sci 8:189 3. Hancock JT, Whiteman M (2014) Hydrogen sulfide and cell signaling: team player or referee? Plant Physiol Biochem 78:37–42

4. Hancock JT, Craig T, Whiteman M (2017) Competition of reactive signals and thiol modifications of proteins. J Cell Signal 2:170 5. Shetty NP, Lyngs Jorgensen HJ, Jensen JD, Collinge DB, Shetty HS (2008) Roles of reactive oxygen species in interactions between plants and pathogens. Eur J Plant Pathol 121:267–280 6. Palmer RM, Ferrige AG, Moncada S (1987) Nitric oxide release accounts for the biological

Addition of Redox Compounds activity of endothelium-derived relaxing factor. Nature 327:524–526 7. Jeandroz S, Wipf D, Stuejr DJ, Lamattina L, Melkonian M, Tian Z, Zhu Y, Carpenter EJ, Wong GK, Wendehenne D (2016) Occurrence, structure, and evolution of nitric oxide synthase-like proteins in the plant kingdom. Sci Signal 9(417):re2 8. Neill S, Desikan R, Hancock JT (2003) Nitric oxide signalling in plants. New Phytol 159:11–35 9. Lisjak M, Teklic T, Wilson ID, Whiteman M, Hancock JT (2013) Hydrogen sulfide: environmental factor or signaling molecule? Plant Cell Environ 36:1607–1616 10. Miller DL, Roth MB (2007) Hydrogen sulfide increases thermotolerance and lifespan in Caenorhabditis elegans. Proc Natl Acad Sci U S A 104:20618–20622 11. Wilson HR, Veal D, Whiteman M, Hancock JT (2017) Hydrogen gas and its role in cell signaling. CAB Rev 12:1–3 12. Li HM, Shen L, Ge JW, Zhang RF (2018) The transfer of hydrogen from inert gas to therapeutic gas. Med Gas Res 7:265–272 13. Zeng J, Ye Z, Sun X (2014) Progress in the study of biological effects of hydrogen on higher plants and its promising application in agriculture. Med Gas Res 4:15 14. Thomas DD, Miranda KM, Espey MG, Citrin D, Jourd’Heuil D, Paolocci N, Hewett SJ, Colton CA, Grisham MB, Feelisch M, Wink DA (2002) Guide for the use of nitric oxide (NO) donors s probes of the chemistry of NO and related redox species in biological systems. Methods Enzymol 359:84–105 15. Li L, Whiteman M, Guan YY, Neo KL, Cheng Y, Lee SW, Zhao Y, Baskar R, Tan CH, Moore PK (2008) Characterization of a novel, water-soluble hydrogen sulfide-releasing molecule (GYY4137): new insights into the biology of hydrogen sulfide. Circulation 117:2351–2360 16. Szczesny B, Modis K, Yanagi K, Coletta C, Le Trionnaire S, Perry A, Wood ME, Whitemna M, Szabo C (2014) AP39, a novel mitochondria-targeted hydrogen sulfide donor, stimulates cellular bioenergetics, exerts cytoprotective effects and protects against the loss of mitochondrial DNA integrity in oxidatively stressed endothelial cells in vitro. Nitric Oxide 41:120–130

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17. Cross AR, Parkinson JF, Jones OT (1984) The superoxide-generating oxidase of leucocytes. NADPH-dependent reduction of flavin and cytochrome b in solubilized preparations. Biochem J 223:337–344 18. Oh H, Siano B, Diamond S (2008) Neutrophil isolation protocol. J Vis Exp 17:745 19. Uy B, McGlashan SR, Shaikh SB (2011) Measurement of reactive oxygen species in the culture media using Acridin Lumingen PS-3 assay. J Biomol Tech 22:95–107 20. Zacharia IG, Deen WM (2005) Diffusivity and solubility of nitric oxide in water and saline. Ann Biomed Eng 33:214–222 21. Bethke PC, Libourell IG, Reinohl V, Jones RL (2006) Sodium nitroprusside, cyanide, nitrite, and nitrate break Arabidopsis seed dormancy in a nitric oxide-dependent manner. Planta 223:805–812 22. Mata-Perez C, Sanchez-Calvo B, Padilla MN, Begara-Morales JC, Valderrama R, Corpas FJ, Barroso JB (2017) Nitro-fatty acids in plant signaling: new key mediators of nitric oxide metabolism. Redox Biol 11:554–561 23. Gupta KJ, Igamberdiev AU (2013) Recommendations of using at least two different methods for measuring NO. Front Plant Sci 4:58 24. Zhao Y, Biggs TD, Xian M (2014) Hydrogen sulfide (H2S) releasing agents: chemistry and biological applications. Chem Commun (Camb) 50:11788–11805 25. Lee JL, Mather AE (1977) Solubility of hydrogen sulfide in water. Ber Bunsenges Phys Chem 81:1020–1023 26. Lisjak M, Teklic T, Wilson ID, Wood ME, Whiteman M, Hancock JT (2011) Hydrogen sulfide effects on stomatal apertures. Plant Signal Behav 6:1444–1446 27. Lee ZW, Zhou J, Chen C-S, Zhao Y, Tan C-H, Li L, Moore PK, Deng L-W (2011) The slowreleasing hydrogen sulfide donor, GYY4137, exhibits novel anti-cancer effects in vitro and in vivo. PLoS One 6:e21077 28. Shen X, Kolluru GK, Yuan S, Kevil CG (2015) Measurement of H2S in vivo and in vitro by the monobromobimane method. Methods Enzymol 554:31–45 29. Renwick GM, Giumarro C, Siegel SM (1964) Hydrogen metabolism in higher plants. Plant Physiol 39:303–306 30. Molecular Hydrogen Foundation: www. molecularhydrogenfoundation.org

Chapter 3 Investigating ROS, RNS, and H2S-Sensitive Signaling Proteins Eleanor Williams, Matthew Whiteman, Mark E. Wood, Ian D. Wilson, Michael R. Ladomery, Joel Allainguillaume, Tihana Teklic, Miro Lisjak, and John T. Hancock Abstract The modification of proteins is a key way to alter their activity and function. Often thiols, cysteine residues, on proteins are attractive targets for such modification. Assuming that the thiol group is accessible then reactions may take place with a range of chemicals found in cells. These may include reactive oxygen species (ROS), such as hydrogen peroxide (H2O2), reactive nitrogen species such as nitric oxide (NO), hydrogen sulfide (H2S), or glutathione. Such modifications often are instrumental to important cellular signaling processes, which ultimately result in modification of physiology of the organism. Therefore, there is a need to be able to identify such modifications. There are a variety of techniques to find proteins which may be altered in this way but here the focus is on two approaches: firstly, the use of fluorescent thiol derivatives and the subsequent use of mass spectrometry to identify the thiols involved; secondly the confirmation of such changes using biochemical assays and genetic mutants. The discussion will be based on the use of two model organisms: firstly the plant Arabidopsis thaliana (both as cell cultures and whole plants) and secondly the nematode worm Caenorhabditis elegans. However, these tools, as described, may be used in a much wider range of biological systems, including human and human tissue cultures. Key words Glutathione, Glyceraldehyde 3-phosphate dehydrogenase, Histidine kinase, Hydrogen peroxide, Hydrogen sulfide, 50 -Iodoacetamide fluorescein, Nitric oxide, Reactive nitrogen species, Reactive oxygen species, Stomatal guard cells, Thiol labeling

1

Introduction Compounds such as reactive oxygen species (ROS), reactive nitrogen species (RNS), hydrogen sulfide or glutathione are known to be involved in the modification of the intracellular redox status [1]. Such compounds are also involved in the modification of thiol residues on proteins. Therefore, such redox signals are an important mechanism by which redox signal transduction occurs in cells. Sensing of redox signals, via their modification of protein

John T. Hancock and Myra E. Conway (eds.), Redox-Mediated Signal Transduction: Methods and Protocols, Methods in Molecular Biology, vol. 1990, https://doi.org/10.1007/978-1-4939-9463-2_3, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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thiols may cause changes in the structure and hence function of the protein. This may then initiate a series of signaling cascades [2]. Therefore, the identification of proteins, which can be modified, is very important for understanding the mechanisms of redox signaling. There are different methods, which may be used for the identification of such proteins. This includes tagging thiol residues with detectable groups such that the tagged protein can be subsequently identified, or, using proteins mutated in thiol residues, either as a pure protein or in its native organism, to analyze its function. Once identification of the protein has been made confirmation, using biochemical techniques is often required. Here examples of such methods and techniques will be presented. 1.1 Outline to Procedure Using Iodoacetamide Compounds to Identify Proteins

The basis of the identification of the reactive thiols in proteins in the presence of the redox compound of choice is to set up a competition reaction. Iodoacetamide can react with thiols and also can be labeled (e.g., with a fluorescent tag). Fluorescent covalently modified iodoacetamide (which can be purchased premade, for example 50 -iodoacetamide fluorescein (50 -IAF)) will react with free thiol groups on proteins. However, when the protein is pre-treated with the redox compound, for example oxidized by ROS, the iodoacetamide group can no longer react with the thiol group as it is no longer in the correct state (see Fig. 1). Therefore, the effect of ROS on thiol modification can be assessed by carrying out experiments in the presence and absence of ROS, along with the iodoacetamide dye, subsequently followed by protein analysis. In the presence of only 50 -IAF any proteins with an available thiol will be rendered fluorescent and be seen as a fluorescent band on subsequent electrophoresis gels. In the presence of ROS such thiols will be oxidized, will no longer be able to react with the 50 -IAF and therefore will not appear on the electrophoresis gel. Therefore, it is the disappearance of the band which is looked for on analysis. This approach can be used not just with ROS but with any compound that may react with the thiol and so render it inaccessible for the reaction with the 50 -IAF. Recently this has been used to identify hydrogen sulfide reactive proteins in the nematode worm C. elegans [3].

Fig. 1 A competition can be set up between the reactive compounds. This can be studies using 50 -IAF, usually causing proteins to fail to gain fluorescence and therefore seemingly to disappear from subsequent electrophoretic analysis

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As an example of using such an approach it was shown that the plant enzyme cytosolic glyceraldehyde 3-phosphate dehydrogenase (GAPDH) is a target of hydrogen peroxide modification [4]. This has also been confirmed by others, and subsequently it has been found that this enzyme, which was originally thought to only be involved in glycolytic pathways, is actually also involved in redox signaling [5]. On thiol modification, it has been found to move to the nucleus where it affects gene expression. Such subsequent data confirms that this relatively simple “fishing trip” for thiol-modified proteins [4] can find interesting and important proteins, which are involved in signaling [5]. Therefore, the first tool described in this chapter is the iodoacetamide labeling procedure followed by proteomics analysis of the labeled proteins. 1.2 Other Methods of Identification of Modified Thiols

Here, a relatively simple method for the identification of modified thiols is described. However, it is not the only method which is available for the purpose. With the interest in the role of NO, it was soon realized that not all the effects of NO were mediated by its effects on metals—such as activation of guanylyl cyclase, which involves the reaction of NO with the heme in the cyclase enzyme. Therefore, there was a requirement to understand how NO and proteins could directly interact together. To this end, a method commonly known as the “biotin switch assay” [6, 7] was developed. Although this method was described over 15 years ago [8], it has been adopted by numerous research groups. The rationale underpinning this assay is different to that described in this chapter. The basis of the biotin switch assay is that it is assumed that thiol groups may have reacted to become S-nitrosated. However, not all of the available thiols would have reacted with NO. Any that are still free thiols are then reacted with methyl-methane thiosulfonate (MMTS) so that at this point all the thiols should be modified, that is, either S-nitrosated or S-methylthionated. The S-nitrosated thiols are then reduced with ascorbate, removing the NO group and so reforming the –SH group. The thiols that have reacted with MMTS are unaffected. It is also assumed that this reaction is specific. It is assumed that thiols which have reacted with glutathione or have been oxidized by ROS are not re-reduced. Subsequently the newly formed –SH groups (those which had previously been nitrosated) are reacted with biotin-HPDP. Hence, they can now be reacted, and labeled, with avidin–horseradish peroxidase. Interestingly, this assay has not been free of criticism, and many researchers say that it needs to be carried out with some caution and care [7, 9]. In the biotin switch assay any S-nitrosated thiol (–SNO) needs to be converted back to the thiol, and then it is subsequently identified. In the assay described in this chapter, there is no such requirement. Furthermore, it is easy to undertake. Previous studies have shown that the assay has been successful in identifying both

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ROS-sensitive proteins [4] and those which may be modified by H2S [3]. Thiols may be sensitive to a range of other compounds, and there is no reason why this method should not be adapted to look for these modifications too. 1.3 The Use of Model Organisms and Mutants

Although techniques and tools described here can be used with any biological material, there is a major advantage in using a model organism. These are usually organisms which are well characterized, both from a biochemical and genetic point of view, and are usually cheap and easy to grow. Much work is carried out on humans but tissues are often hard to source. Therefore, carrying out experiments on other organisms, with the data subsequently informing human studies, is often a sensible way forward. Here two model organisms are used. For plant science Arabidopsis thaliana, is used and for animal science, the nematode Caenorhabditis elegans is used. Others have extensively used both. More importantly they have genomes that have been well characterized and for both mutants are available. For information on C. elegans see [10] while for a review on Arabidopsis see [11]. Sydney Brenner suggested, in 1965, the use of the small nematode C. elegans [12]. It is very cheap to grow and easy to maintain either on plates or in liquid culture. It also has a short life span, so it can be studied in many developmental phases easily. Importantly, its developmental stages, along with its genome, have been very well characterized. Each worm has exactly 959 somatic cells, and the development and characteristics of those cells are well studied. The homology between genes in C. elegans and those in humans also makes it an attractive model, and they are a superb resource for the study of human diseases and for drug discovery [13]. Redox biology has not ignored C. elegans. Interestingly treatment of C. elegans with H2S increases their lifespan and bestows on the worms thermal tolerance [14]. H2S has been suggested to be an important compound, which may protect organisms from disease. Therefore using C. elegans to understand the roles and interactions of redox compounds [15] would help to unravel the redox biology in a range of organisms, including humans. As with C. elegans, there is a huge international community and a wealth of knowledge on the model plant Arabidopsis thaliana. This has also led to development of various public resources, such as the availability of seed stocks of different mutants via stock centres. As will be discussed below this allows the use of mutants either to search for redox sensitive proteins or to confirm the identification of such proteins using the iodoacetamide tagging approach.

1.4 The Treatment of Organisms with ROS, RNS or H2S

The most likely member of the ROS family of compounds, which is going to be used for these types of experiments, is hydrogen peroxide (H2O2) and this can be readily purchased from regular suppliers. Other ROS, which may be considered to be used, include

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the superoxide anion, which will dismute to H2O2 very readily. Superoxide can be purchased as a salt, usually potassium superoxide. On the other hand, the addition of gases such as NO and H2S are more problematic, both can be purchased in gaseous form and it is possible to either create saturated solutions of the gas, which can then be applied, or the biological material can be placed in the presence of the gas in either an air or nitrogen atmosphere. Such experiments may be applicable to plant material for example. However, using gasses such as NO and H2S are not without problems and in fact risk (see Note 1). H2S is a gas. Therefore, by using gas tanks of H2S (these can be purchased, for example, from Praxair) samples can be treated directly with H2S, that is, from the gaseous phase. This may be a good option if whole organisms are to be treated, or parts of an organism, such as the leaves of the plant. Even so, strict safety measures will be needed due to the toxicity of H2S—any work with H2S needs to be well ventilated. However, it needs to be recognized that the use of gas is not always convenient, or indeed suitable for treating biological samples. This may include work being carried out on cells and cell cultures, or protein samples. To this end, compounds which release H2S are used (these are often referred to as donor molecules.) The most common donors include sodium hydrosulfide (NaHS) and sodium sulfide (Na2S). These are cheap and easy to obtain. They are also easy to store and use. A good example of such use is the study which investigated the mitogen-activated protein kinase-mediated apoptosis of animal cells [16]. However, there needs to be a word of caution here. As soon as these compounds are in solution they will release H2S quickly. Samples will therefore be exposed to relatively high H2S for a short period of time. To overcome this new donor compounds have been designed. One of these compounds is known as GYY4137 [17]. Once in solution this compound will release H2S in a more physiological manner, that is, more slowly for longer periods. Development of similar compounds has led to some which are able to target the release of H2S to organelles. A range of such compounds are now becoming commercially available (e.g. from VIVA Biosciences), and it is thought that in the future compounds such as these might be the basis for a new therapeutic strategy for a range of diseases [18].

2

Materials 1. AT3 medium: Murashige and Skoog medium supplemented with 3% sucrose, 0.5 mg/L NAA, and 0.05 mg/L kinetin, pH 5.5. 2. Murashige and Skoog media.

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3. Extraction buffer: 50 mM Tris–HCl pH 7.5, with complete protease inhibitor cocktail. 4. Equilibration solution: 50 mM Tris–HCl, pH 8.8, 6 M urea, 30% glycerol, 2% SDS, 0.025% bromophenol blue. 5. Protease inhibitor cocktail tablets. 6. 50 -Iodoacetamide fluorescein (50 -IAF) made as a stock solution fresh (for example: 10 mM stock in DMSO). 7. 2D electrophoresis solutions and reagents: all PlusOne reagents. 8. H2O2 solution (30% v/v). 9. Stomatal bioassay buffer: 10 mM MES. 5 mM KCl, 100 μM CaCl2, pH 6.15 with 1 M Tris–HCl, pH 7.5. 10. Levington’s F2 compost with sand. 11. Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) assay buffer: 100 mM Tris–HCl, pH 8, 10 mM MgCl2, 0.2 mM NADH, 5 mM ATP, 2 mM phosphoglyceric acid (PGA). 12. Isolated GAPDH isolated enzyme, for example from rabbit muscle. 13. MES/KCl buffer: 5 mM KCl, 10 mM Mes, 50 μM CaCl2, pH 6.15. 14. Seed sterilizing solution: 10% bleach, 0.1% SDS. 15. H2S donor molecules such as NaHS, Na2S, or GYY4137.

3

Methods

3.1 Growth and Maintenance of Arabidopsis thaliana Cell Suspension Cultures

Suspension cultures of Arabidopsis thaliana (obtained from an established source) must be maintained under aseptic conditions as follows: 1. Once a week, transfer 10 mL of a 7-day culture into 100 mL of AT3 medium in sterile conical flasks (see Note 2). This should be performed in a laminar flow cabinet. 2. Maintain the cells in a controlled growth environment room/ chamber at 24  C in the light (16 h, 100–150 μE/m2/s) on a rotary shaker at 110 rpm. 3. When cell cultures are required for treatment, aliquot out the required volume of cells (note their packed cell volume) into sterile Falcon tubes or equivalent. Following treatment, harvest the cells by vacuum filtration through Whatman paper and a Buchner funnel. Cells are subsequently frozen in liquid nitrogen and stored at 80  C until further use.

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3.2 Growth and Maintenance of C. elegans

3.3 Treatment of Cells and Labeling of Proteins

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C. elegans can be grown on NGM plates (Nematode Growth Media plates) or they can be grown in liquid culture [10] (see Note 3). Usually they will grow with E. coli OP50 as a food source. Often enough worms can be sourced from plates, making liquid culture unnecessary. For NGM plates: mix 3 g NaCl, 17 g agar, 2.5 g peptone and 975 mL H2O. This should then be autoclaved and cooled. To this add: 1 mL of 1 M CaCl2; 1 mL of 5 mg/mL cholesterol in ethanol; 1 mL of 1 M MgSO4; 25 mL of 1 M KPO4 buffer. Although any size plates can be used, ideal for the growth of the worms are large square petri dishes (120  120  17 mm). Once the plates are poured they need to be spread with E. coli using a glass rod. The plates then need to be incubated at 37  C for at least 6 h. Alternatively, they can be incubated overnight at room temperature. Following incubation, ensure that the plates are cooled to room temperature before the worms are added. Basically, worms are added by a method called ‘chunking’. To do these chunks of NGM plate containing worms and E. coli are removed from one plate and added to a new plate. This allows the worms to multiply on the new food source. Alternatively, worms and E. coli can be washed from a well-populated plate. The resultant liquid needs to be briefly centrifuged at 1500  g. The majority of the supernatant should be removed and then some of the supernatant remaining in the sample can be pipetted onto a fresh E. coli containing plate. E. coli ideally should be grown at 37  C but a 20  C incubator should be used for growing the worms. Worms should be placed onto new plates every few days to prevent over population of the plates and restriction of their source of food. 1. For treatment with H2O2, cell cultures of equal cell densities are aliquoted into sterile Falcon tubes, or equivalent. Concentrations of H2O2 that are physiological to the system being used (in this case, cell cultures) are applied to the cell cultures and left for a defined time period. For example, apply 10 mM H2O2 to cell cultures of a known volume, for 10 min with constant shaking (see Note 4). Once the treatment is complete, harvest the cells by filtration as described above: step 3 of Subheading 3.1. 2. Extract proteins from harvested cells as follows. Grind the filtered cell cultures in a prechilled mortar and pestle at 4  C in extraction buffer (see Note 5). 3. Centrifuge the homogenate in microfuge tubes at 12,000  g, 10 min. 4. Remove the supernatant and recentrifuge this supernatant in a fresh tube at 12,000  g, 10 min.

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5. Determine protein concentrations using a reliable protein determination assay (e.g., Bradford assay or equivalent). 6. For labeling the proteins, use equal concentrations of proteins across the treatment range, in a defined volume, that is, adjust the volume of the samples so that they are equal. To each protein extract, add 50 -IAF to a final concentration of 100 μM (from a stock solution of 50 -IAF of 10 mM in DMSO) and incubate in the dark at room temperature for 10 min. 3.4 Analysis of Proteins Using Electrophoresis

Proteins labeled as above can be analyzed by either one or two-dimensional gel electrophoresis. For one dimensional gel electrophoresis (1D SDS), use standard procedures as used for SDS-PAGE, using a 12% resolving gel, and in the dark to avoid photobleaching of the dye. Once completely resolved, scan the gels using a Typhoon scanner at an excitation wavelength of 490–495 nm and emission wavelength of 515–520 nm. For identification of proteins, subsequently stain the gels with colloidal Coomassie as per the manufacturer’s guidelines. For two-dimensional gel electrophoresis (2D gel electrophoresis), different procedures are followed, which involves accurate quantification followed by electrophoresis according to the pI and subsequently the MW of the proteins being analyzed. Following electrophoresis, labeled proteins can be visualized (Typhoon scanner at an excitation wavelength of 490–495 nm and emission wavelength of 515–520 nm) and identification of proteins performed by mass spectrometry. The procedures are described briefly below; however, methods differ between different manufacturers. The procedure described here is according to that prescribed by GE Healthcare, UK, for 2D gel electrophoresis. 1. Extract the proteins Subheading 3.2.

as

described

in

steps

1–4

of

2. Determine the concentration of soluble proteins using a 2D Quantification kit (see Note 6). 3. Add 100 μM 50 -IAF to the protein extracts (of equal concentration, 50 μg) for 10 min at room temperature. 4. Precipitate the extracts with TCA/acetone (see Note 7) and resuspend the pellet in rehydration buffer (8 M urea, 2% CHAPS). 5. Add 0.5% IPG buffer and 2.8 mg/mL DTT to the samples, and analyze by electrophoresis as follows: IEF overnight on 24 cm pH 3–10 nonlinear Immobiline DryStrip gels using an IPGPhor unit, in the dark, with the following settings: 500 V for 10 min, 4000 V for 1 h 30 min, 8000 V for 6 h 30 min at 20  C. At this point, the strips are ready to be electrophoresed on the second dimension.

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6. For the second dimension, strips are first equilibrated for 15 min in equilibration solution containing 0.5% (w/v) DTT and then for 15 min in equilibration solution containing 4.5% (w/v) iodoacetamide for 15 min. 7. Resolve the proteins by loading the strips onto 12.5% SDS gels prepared using the Ettan DALTsix gel system at 20  C at an initial power setting of 2.5 W per gel for 30 min, followed by 18 W per gel for 5 h. 8. Following electrophoresis scan the gels on the Typhoon scanner as described above (Subheading 3.3). 9. Following scanning, stain the gels using colloidal Coomassie stain according to the manufacturer’s guidelines. 3.5 Identification of Proteins Using MALDI-ToF

Following electrophoresis, any spots or bands identified as being labeled with 50 -IAF has to be identified. For this, the protein band or spot is excised using a sharp scalpel blade from the acrylamide gel and under very clean conditions, placed in a 1.5 mL tube. The procedure is repeated for every spot or band identified as being differentially labeled on the gel. Samples are subsequently digested using trypsin. 1. Chop the spot/band into small (1 mm  1 mm) pieces; this is best done in the tube using a clean scalpel or needle. 2. Wash the gel pieces in 150 μL water for 5 min. Centrifuge for 3 min and remove supernatant. 3. Add 150 μL acetonitrile and leave for 10–15 min to dehydrate the gel pieces. 4. Remove all acetonitrile and add approximately 50 μL 10 mM DTT in 0.1 mM NH4HCO3. Incubate for 30 min at 56  C. 5. Remove DTT and add acetonitrile to dehydrate the gel pieces as before. 6. Centrifuge for 3 min and remove supernatant and add 50 μL 55 mM iodoacetamide in 0.1 mM NH4HCO3 and leave in the dark for 45 min. 7. Centrifuge for 3 min and remove supernatant and wash with 150 μL 0.1 mM NH4HCO3 for 15 min. 8. Centrifuge for 3 min and remove supernatant and add acetonitrile to dehydrate the gel pieces as before. 9. Dry the gel pieces in a vacuum centrifuge. 10. Prepare a trypsin solution in 50 mM NH4HCO3 containing 12.5 ng/μL Trypsin (see Note 8). 11. Add 20 μL of this trypsin solution and incubate at 4  C for 30–45 min. Check after 15–20 min and top up with more trypsin solution if all liquid has been absorbed by the gel piece.

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12. After 30–45 min remove all remaining liquid and add 20 μL of 50 mM NH4HCO3 without trypsin. 13. Incubate at 37  C for 16 h. 14. To extract the peptides, add 10 μL 50% acetonitrile/5% formic acid and place tube in a sonicating water bath for 10 min. 15. Remove liquid and transfer to a clean tube (this should contain the peptides). 16. Repeat the sonication once more using 10–20 μL 50% acetonitrile–5% formic acid, each time transferring the liquid to the clean tube. 17. Mix an aliquot of the extracted peptides (5 μL) with 5 μL matrix solution. 18. Analyze an aliquot of the sample (1 μL) by MALDI-ToF mass spectrometry. 19. To confirm the identity of proteins, peptides can also be analyzed using LC-ESI-MS/MS, using a C18 Pepmap column, with a 5–80% acetonitrile gradient with 0.1% formic acid, over 60 min with a flow rate of 200 nL/min. Run the ESI-MS/MS in data dependent acquisition (DDA) mode with a capillary voltage of 3.5 kV. 20. Bioinformatic analyses should be carried out using ProteinLynx Global Server. Using the above procedures, GAPDH (Accession number: P25858) was identified, from Arabidopsis suspension cultures, as being labeled with 50 -IAF, and in the presence of H2O2 the fluorescence intensity of this protein decreased significantly. Based on these findings it was concluded that GAPDH in Arabidopsis cell cultures is a target of H2O2 signaling [4]. In order to prove a functional role for this, it is important to ascertain whether or not redox modification would actually change the function of the protein that has been labeled. To test this, assays for the activity of GAPDH need to be performed, and the response to H2O2 tested. 3.6 Assays for GAPDH Activity

In order to determine if H2O2 does affect the function of the protein GAPDH identified above using thiol-tagging and proteomics, activity of the enzyme GAPDH is tested in the absence and presence of H2O2. For this either pure commercial enzyme or plant extract (soluble proteins, as obtained in steps 1–4 of Subheading 3.2) can be used. GAPDH activity can be determined using a coupled NADHdependent reaction. The reactions catalyzed are as follows: ðPPK; MgCl2 Þ

PGA þ ATP ! 1,3‐bisphosphoglycerate þ ADP

ð1Þ

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37

1,3‐Bisphosphoglycerate þ NADH ðGAPDHÞ

þ Hþ ! glyceraldehyde 3‐phosphate þ NADþ þ Pi ð2Þ

1. Into a cuvette place 3 mL GAPDH assay buffer (see Subheading 2). 2. Add either 35 units of the isolated enzyme or soluble protein extract from plants (see Note 7). 3. Start the reaction by adding three units of phosphoglyceric phosphokinase (PPK), and monitor the reduction in absorbance following NADH oxidation at 340 nm for 10 min using a spectrophotometer linked to a chart recorder. 4. For monitoring the effect of H2O2 on this activity, add different concentrations of H2O2 to the reaction mixture for 5 min, followed by the addition of PPK. Monitor the activity for 10 min (see Note 9). 5. Repeat the experiment at least three times with either different batches of enzyme or plant extract. 6. In order to show that the effect of H2O2 is actually reversible by using a reducing agent, reversal of inhibition of GAPDH activity is performed by starting the reaction with PPK, adding H2O2 for 10 min, following which glutathione (10 mM) or DTT (10 mM) is added and activity continued to be monitored (see Note 10). In order to confirm that the activity from the extract is indeed from the GAPDH protein identified via proteomics, molecular biology experiments need to be performed, which will involve the cloning and expression of the GAPDH protein, followed by purification of the pure protein. Subsequently, experiments such as above can be performed on this pure plant protein. In addition to identify the exact post-translational modification that may have occurred on the protein by H2O2, use mass spectrometry. 3.7 Growth and Analysis of Arabidopsis Plants

To study the effect of Cys-containing mutations (or any mutation) in plant proteins, one can make use of the different resources available for such purposes within the Arabidopsis plant community. A number of researchers have donated seed stocks of Arabidopsis to a national seed stock center, from where, for a small price, seeds can be obtained for noncommercial purposes by any researcher (see http://www.nasc.org). With a knowledge of what plant is needed, the holdings of the stock center can be searched for various seeds that have been donated. Using this approach, a family of genes called histidine kinases were screened for testing the ability of these proteins to act as ROS

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sensors. One group of proteins in this family belongs to the class of ethylene receptors. Ethylene is a plant hormone involved in a number of physiological processes such as fruit ripening, falling of leaves and formation of root hairs. A group of five receptors of the histidine kinase family act as ethylene receptors. A large amount of research into ethylene signaling has involved the use of Arabidopsis plants mutated in these receptors. One such mutant is mutated in a Cys residue at the ethylene binding site. Therefore the role of this Cys, and hence this protein, in ROS signaling in plants has been explored by studying different physiological responses to ROS [19]. Some of these include the effect of hydrogen peroxide on stomatal closure, gene expression and root growth. By comparison of wild type and mutant plants grown under exactly the same conditions, analysis of the requirement for a functional protein in ROS signaling can be achieved. For example, the ethylene receptor ETR1 has a number of mutants that are available. Of this, the etr1–1 mutant has a mutation in a particular cysteine residue, Cys65, which is required for ethylene binding. By hypothesizing that this Cys65 might be essential for ROS sensing and signaling, various experiments can be designed to test this parameter. Seeds of etr1–1 mutant plants were obtained from the stock center. Simultaneously, the corresponding wild type background plant seeds were also obtained. 3.7.1 Growth of Arabidopsis Plants

1. When the seeds have been obtained, store them at 4  C for at least 48 h, to break the dormancy of the seeds (see Note 11). 2. Sow both wild type and mutant etr1–1 (mutant) seeds in separate trays of Levington’s F2 compost with sand, after wetting the compost until it formed clumps in the hand (see Note 12). Sow the seeds evenly spaced out, and singly, using a toothpick with a wet end. On average, approximately 15–18 plants can be sown on a 30 cm  20 cm tray. 3. Place the trays in a controlled environment growth cabinet with the following settings: 22  C, 80% relative humidity, and 16 h photoperiod with lights of intensity 60–100 μE m2 s1 (see Note 13). 4. Water the trays from below until the bottom of the tray is just above covered level (see Note 14). Water the plants three times a week, Monday, Wednesday and Friday at similar times, to maintain uniformity in plant growth. 5. Let the plants grow for at least 4 weeks until fully expanded rosette leaves have been obtained. Use leaves of similar size from both wild type and mutant plants.

S-Nitrosation and S-Glutathionylation 3.7.2 Phenotypic Analysis of Plants Stomatal Bioassays

39

H2O2 causes stomatal closure in wild type Arabidopsis leaves [19] and this can easily be used as a system to study the function of various signaling proteins. In order to perform stomatal bioassays, it is of absolute importance that both wild-type and mutant plants are grown under exactly the same conditions, preferably in the same growth cabinet. This avoids any differences and variations due to growth conditions. 1. Detach single leaves from different plants that have been grown for 4 weeks (see Note 15). 2. Place each individual detached leaf in a 5 mL Petri dish with 2 mL of MES/KCl buffer. 3. Cover the dishes and place them in the growth cabinet for 2.5 h. 4. Add various doses of H2O2 to different Petri dishes (wild-type and mutant leaves) (see Note 16). 5. Incubate for a further 2.5 h under the same conditions. 6. Blend each leaf individually in a Waring blender with 100 mL of water, for 30 s. 7. Collect the blended epidermal fragments on a 100 μm nylon mesh. 8. Transfer the epidermal fragments to a glass microscope slide and place a cover slip on top. 9. Measure the stomatal apertures with the aid of an eyepiece graticule on a calibrated microscope. During measurements, make sure to move around and note apertures of at least 25 different stomata, from different fields of view. Also, ensure that the correct focal plane for the aperture is used. As the cell walls of guard cells are fairly thick, it can be difficult to focus on the inner walls of the aperture (see Fig. 2). Also, it is very

Fig. 2 Light microscope image of an open Arabidopsis stomate. Each cell surrounding the stomate is a guard cell. The arrow indicates the stomatal aperture (width) which is measured

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important to repeat these experiments at least 3–5 times preferably with different batches of plants. Using this method, the results from mutant plants can be obtained [19]. In that study with increasing concentrations of exogenous H2O2, stomata of wild-type guard cells respond by closing, whilst stomata of the mutant etr1–1 plants do not close in response to H2O2. As the etr1–1 mutant is mutated in the Cys65 residue this confirms that this residue is somehow involved in mediating H2O2-induced closure in Arabidopsis guard cells.

4

Notes 1. When handling gases precautions should be taken. As well as writing and understanding appropriate rick assessments usually such experiments need to be carried out in fume hoods. Both NO and H2S are toxic and can be fatal. 2. Sterilize flasks by baking at 200  C for 2 h. 3. Although C. elegans is a lower animal it still needs to be considered as an animal and some institutes will require its use to be agreed by an Animal Welfare Committee, or equivalent. 4. Time of H2O2 treatment is determined empirically for different tissue types. The time mentioned here has been tested for cell cultures. 5. Volume of buffer to sample: 1.5 mL buffer: 3 g fresh weight tissue, calculated by aspirating all media from cell cultures and weighing wet weight of cells in Falcon tube. 6. Soluble protein extract is obtained by homogenizing the cells, followed by centrifugation at 12,000  g for 10 min to obtain soluble proteins. 7. For TCA/acetone precipitation, add ice-cold 100% TCA to make a final concentration of 10% (v/v). Chill on ice for 15 min and centrifuge the sample for 20 min to remove the supernatant. Wash the pellet with 1 mL of 100% acetone, vortex well and recentrifuge for 10 min. Repeat this process twice to wash the pellet. Air-dry the pellet and resuspend in the required buffer. 8. If for some reason the protein does not digest into peptides easily using trypsin, then alternative enzymes can be used (e.g., chymotrypsin). 9. The activity of GAPDH can be followed by the oxidation of NADH as it is a stoichiometric reaction.

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10. To confirm that the inhibition of GAPDH is by effects on the enzyme itself, rather than on the phosphoglyceric phosphokinase (PPK), the assay can be carried out in the reverse direction, using GAPDH as the substrate and following the reduction of NADH instead of its oxidation. 11. The amount of seeds in a microfuge tube which comes up to the 0.1 mL mark is approximately 300–500 seeds. 12. Overwetting should be avoided. 13. It is very important to have the lights of the appropriate intensity as this vastly affects the growth of the plants. 14. Overwatering of plants must be avoided. 15. If the plants start flowering do not use them for the stomatal assays. 16. For getting accurate concentrations of H2O2 in each plate, make a 1000 stock of the required concentration of H2O2, and dilute this in 50 mL of medium in Falcon tubes, enough to pour two plates. Repeat with fresh tubes for each dose. References 1. Hancock JT (2009) The role of redox mechanisms in cell signalling. Mol Biotechnol 43:162–166 2. Cooper C, Patel RP, Brookes PS, DarleyUsmar VM (2002) Nanotransducers in cellular redox signalling: modification of thiols by reactive oxygen and reactive nitrogen species. Trends Biochem Sci 27:489–492 3. Williams E, Pead S, Whiteman M, Wood ME, Wilson ID, Ladomery MR, Teklic T, Lisjak M, Hancock JT (2015) Detection of thiol modifications by hydrogen sulfide. Methods Enzymol 555:233–251 4. Hancock JT, Henson D, Nyirenda M, Desikan R, Harrison J, Lewis M, Hughes J, Neill SJ (2005) Proteomic identification of glyceraldehyde 3-phosphate dehydrogenase as an inhibitory target of hydrogen peroxide in Arabidopsis. Plant Physiol Biochem 43:828–835 5. Yang SS, Zhai QH (2017) Cytosolic GAPDH: a key mediator in redox signal transduction in plants. Biol Plant 61:417–426. https://doi. org/10.1007/s10535-017-0706-y 6. Zhang Y, Keszler A, Broniowska KA, Hogg N (2005) Characterization and application of the biotin-switch assay for the identification of S-nitrosated proteins. Free Radic Biol Med 38:871–881 7. Forrester MT, Foster MW, Benhar M, Stamler JS (2009) Detection of protein S-nitrosylation

with the biotin-switch technique. Free Radic Biol Med 46:119–126 8. Jaffrey SR, Erdjument-Bromage H, Ferris CD, Tempst P, Snyder SH (2001) Protein S-nitrosylation: a physiological signal for neuronal nitric oxide. Nat Cell Biol 3:193–197 9. Kovacs I, Lindermayr C (2013) Nitric oxidebased protein modification: formation and sitespecificity of protein S-nitrosylation. Front Plant Sci 4:137 10. WormBook http://www.wormbook.org/ 11. Meinke DW, Cherry JM, Dean C, Rounsley SD, Koornneef M (1998) Arabidopsis thaliana: A model plant for genome analysis. Science 282:662–682 12. Brenner S (1974) The genetics of Caenorhabditis elegans. Genetics 77:71–94 13. Kaletta T, Hengartner MO (2006) Finding function in novel targets: C. elegans as a model organism. Nat Rev Drug Discov 5:387–399 14. Miller DL, Roth MB (2007) Hydrogen sulfide increases thermotolerance and lifespan in Caenorhabditis elegans. Proc Natl Acad Sci U S A 104:20618–20622 15. Hancock JT, Whiteman M (2014) Hydrogen sulfide and cell signaling: team player or referee? Plant Physiol Biochem 78:37–42 16. Adhikari S, Bhatia M (2008) H2S-induced pancreatic acinar cell apoptosis is mediated via

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JNK and p38 MAP kinase. J Cell Mol Med 12:1374–1383 17. Li L, Whiteman M, Guan YY, Neo KL, Cheng Y, Lee SW, Zhao Y, Baskar R, Tan CH, Moore PK (2008) Characterization of a novel, water-soluble hydrogen sulfide-releasing molecule (GYY4137): new insights into the biology of hydrogen sulfide. Circulation 117:2351–2360 18. Szczesny B, Modis K, Yanagi K, Coletta C, Le Trionnaire S, Perry A, Wood ME,

Whitemna M, Szabo C (2014) AP39, a novel mitochondria-targeted hydrogen sulfide donor, stimulates cellular bioenergetics, exerts cytoprotective effects and protects against the loss of mitochondrial DNA integrity in oxidatively stressed endothelial cells in vitro. Nitric Oxide 41:120–130 19. Desikan R, Hancock JT, Bright J, Harrison J, Weir I, Hooley R, Neill SJ (2005) A novel role for ETR1: hydrogen peroxide signalling in stomatal guard cells. Plant Physiol 137:831–834

Chapter 4 Measurement of 4-Hydroxynonenal (4-HNE) Protein Adducts by ELISA Kosha Mehta and Vinood B. Patel Abstract Enzyme linked immunosorbent assay (ELISA) is a widely used technique for the measurement of antigens and antibodies alike. We describe here procedures, indirect ELISA and sandwich ELISA for the detection of 4-hydroxynonenal protein adducts. These adducts are stable compounds formed within cells and bodily fluids under conditions of oxidative stress. They can act as sensitive biomarkers of oxidative stress and are directly linked to disease pathology. Key words ELISA, 4-Hydroxynonenal, 4-HNE, Anti-HNE antibodies, HRP-labeled antibodies, Dot blot, Indirect ELISA, Sandwich ELISA

1

Introduction 4-Hydroxynonenal (4-HNE) is an aldehyde end product of lipid peroxidation. This highly reactive electrophile binds avidly to proteins with a cysteine, histidine or lysine residues, forming stable protein adducts [1]. 4-HNE-protein adducts are elevated in conditions involving oxidative stress [2–4], suggesting they can be a useful biomarker of disease pathology, but also due to its’ immunogenic properties, may directly lead to disease pathogenesis [5]. Protein modification can also directly alter a protein’s function [6]. In addition, 4-HNE has been implicated as a signaling molecule in apoptosis, and through binding of glutathione may affect the cellular redox state [7]. Therefore, measurement of 4-HNE protein adducts may indicate general oxidative stress and/or disease pathology. Enzyme linked immunosorbent assay (ELISA) involves colorimetric or fluorimetric detection and measurement of proteins or antibodies of interest in an unknown sample and is the most commonly used biochemical assay in diagnostic laboratories. It can be classified into various types and subtypes such as competitive,

John T. Hancock and Myra E. Conway (eds.), Redox-Mediated Signal Transduction: Methods and Protocols, Methods in Molecular Biology, vol. 1990, https://doi.org/10.1007/978-1-4939-9463-2_4, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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non-competitive, direct, indirect, displacement and sandwich ELISA; the type is chosen based on the application and specificity required. While the protocols differ slightly, the main principle of ELISA remains the same across all types, i.e. binding of either the antigen (sample/protein) or the antibody to a solid surface and measurement of the intensity of the interaction between the antigen and the antibody. In the following method, we describe indirect and sandwich ELISA using the colorimetric horseradish peroxidase (HRP) system for the measurement of HNE-protein adducts. It is important to note that while the indirect format measures total HNE in a sample (i.e. general oxidative stress), the sandwich format is more specific and detects HNE conjugated to a specific protein of interest. In the indirect ELISA, the unknown sample (or HNE-conjugated protein standard) is absorbed on the surface of the ELISA plate. This is incubated with an anti-HNE antibody (referred as primary antibody) that binds to the HNE moiety on the sample. Further, a HRP-labeled detection antibody (referred as the secondary antibody) is added, which binds to the primary antibody (i.e., the anti-HNE antibody). Subsequent addition of the HRP-specific substrate tetramethylbenzidine (TMB) to this complex allows formation of a yellow-colored product that elicits a chromogenic signal, which is read by the microplate reader (Fig. 1a). In the sandwich format, a “sandwich” is created with at least two antibodies that bind to different and distal epitopes on the sample.

A

B

Key to figure: Well of a 96-well ELISA plate

TMB

Y Y Indirect ELISA

Sample/HNEconjugated protein

Y

Yellow product

Y

Y

Anti-HNE antibody

Y

44

HRP-labelled detection antibody

Y

Capture antibody, specific for the protein of interest

Sandwich ELISA

Fig. 1 Schematic representation of ELISA. A schematic representation of indirect ELISA (a) and sandwich ELISA (b) has been shown. Addition of the HRP-specific substrate TMB produces a yellow color, the intensity of which is measured by the ELISA plate reader. The sandwich ELISA involves binding of two antibodies to different epitopes on the sample, where the capture antibody is specific for the protein of interest. Thus, it is more specific than the indirect ELISA

HNE-Protein Adduct Measurement by ELISA B

Absorbance at 450 nm

1.2 1 0.8

R² = 0.9967

0.6 0.4

0.2 0 0

1 2 3 4 HNE-protein standard (µg/mL)

5

Absorbance at 450 nm

A

45

0.6 0.5 0.4 R² = 0.9976

0.3 0.2 0.1 0 0

1 2 3 4 HNE-protein standard (µg/mL)

5

Fig. 2 Typical ELISA standard curves. Typical examples of the standard curves obtained following an indirect ELISA (a) and a sandwich ELISA (b) are shown. The average absorbance reading of each concentration of the HNE-protein standard is subtracted from the reading of the blank (without standard) and plotted on a graph. The concentration of HNE in unknown samples is interpolated from the graph by using the equation of the line of regression

Typically, one antibody is for capture of the protein of interest and the second antibody is an anti-HNE antibody that is HRP-labeled to facilitate detection. First, the protein-specific antibody is absorbed on the surface of the ELISA plate (referred as the capture antibody) to which the unknown sample or a known concentration of HNE-conjugated protein standard is added. Following this, HRP-labeled anti-HNE detection antibody is added, which binds to a different distal epitope on the sample/standard. Similar to the indirect ELISA, addition of TMB mediates the formation of a yellow-colored product, which is detected by the plate reader and the absorbance is recorded (Fig. 1b). In both indirect and sandwich ELISA, the presence of HNE-protein adducts in the sample is confirmed by comparing the readings of the sample with those of the HNE-protein standard curve (Fig. 2).

2

Materials

2.1 Preparation of Antibodies

1. Antibodies: Procure antibodies for the indirect/sandwich ELISA (see Notes 1 and 2). 2. Anti-HNE antibodies (see Note 2). 3. HRP-labeled detection antibodies for the indirect ELISA (see Notes 3 and 4). 4. Capture antibodies for sandwich ELISA: Polyclonal or monoclonal antibodies are produced against your protein of interest (antigen) by immunizing the desired species using highly purified (>99%) protein. 5. For the sandwich ELISA the generated anti-HNE antibodies (see Note 1) require labeling with HRP, which can be carried out by AbD Serotec (Bio-Rad) or Abcam.

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2.2 ELISA Assay Components

1. Protein standards: HNE-conjugated protein standards are commercially available or can be prepared ‘in house’ [8] (see Note 5). 2. ELISA plates: 96-well, High Bind plates suitable for ELSA application (see Note 6). 3. Disposable plate sealers. 4. Phosphate buffered saline (PBS). 5. Coating buffer: 0.35 M sodium bicarbonate, 0.15 M sodium carbonate (pH 9.6) in deionized water. 6. Wash buffer A: 0.05% PBS-Tween (T) (v/v) (see Note 7). 7. Wash buffer B: 0.1% PBS-T (v/v) (see Note 7). 8. Blocking buffer: Casein (see Note 8). 9. Substrate solution: TMB Substrate solution. 10. Stop solution: 2 M sulfuric acid. 11. Reagent reservoirs, timer, a well-calibrated multichannel pipette and microplate reader. 12. West Pico chemiluminescent kit.

3

Methods All antibodies should be stored at 4  C or 20  C, as suggested by the manufacturer (see Note 2). All buffers and antibody preparations should be equilibrated at room temperature (RT) before addition to the wells and all procedures should be carried out at RT, unless stated otherwise. Buffers for coating, blocking and washing are commercially available or can be prepared “in house” in deionized Milli-Q grade water using analytical grade reagents and stored at RT.

3.1 Optimization of Antibodies, Standards, and Sample

Prior to performing the ELISA with samples, it is important to confirm the binding of the anti-HNE antibody to an HNE-labeled protein and binding of the capture antibody to the ELISA plate. In addition, it is important to determine the optimal concentrations of antibodies and the optimal range of standards for the ELISA. 1. To confirm that the commercially produced anti-HNE antibody binds to a HNE-labeled protein, the dot blot method can be used. Briefly, a known amount (e.g., 30 μg) of a HNE-conjugated protein (i.e., a standard can be used) is blotted on a nitrocellulose membrane, allowed to dry for 10 min and probed with selected concentrations of anti-HNE antibody made in wash buffer A (see Note 9). Following washing with buffer B, the complex is detected by a species compatible HRP-labeled detection antibody by using the West PICO

HNE-Protein Adduct Measurement by ELISA

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HRP-conjugated detection antibody (µg/mL) HNE-protein standards (µg/mL)

0

5

10

15

0

5

10

15

0

5

10

15

0 0.05 0.5 1 2 3 4 5

Capture antibody 5 µg/mL

Capture antibody 10 µg/mL

Capture antibody 15 µg/mL

Fig. 3 Representative titrations of antibodies for sandwich ELISA. The figure is a representative schematic of a 96-well ELISA Plate. It shows combinations of different antibody concentrations probed with a range of known HNE-protein standards to optimize the sandwich ELISA

chemiluminescent kit, as suggested by the manufacturer (see Note 9). 2. For the sandwich ELISA, it is important to confirm first that the capture antibody binds to the ELISA plate. Here the wells are coated with 5 μg/mL and 15 μg/mL of capture antibody, followed by detection using a species compatible HRP-labeled detection antibody, typically between 0.01 μg/mL and 0.04 μg/mL. For this assay, the protocol mentioned in Subheading 3.4, should be followed. 3. To optimize the antibody-concentrations and the concentration range of standards, titrations of various combinations of antibody-concentrations should be performed with a wide range of HNE-conjugated protein standards. An example of the titration board for the sandwich format is suggested in Fig. 3. Following this, the most linear range of the concentration of standards is chosen (Fig. 2). 4. To optimize the sample concentration, different dilutions of the sample in PBS can be used, for example neat sample, 1:1, 1:10, and 1:100. 3.2 Selecting Control Samples

The following experimental controls are added in triplicate to each assay plate. 1. Positive control: A previously assessed sample that showed a positive reaction or a known concentration of HNE-protein that shows a positive result.

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2. Negative controls for indirect ELISA: (a) Blank-1: wells with the known HNE-conjugated protein standard and anti-HNE antibody, but without HRP-labeled detection antibody. Use wash buffer B instead of the detection antibody. (b) Blank-2: wells without HNE-conjugated protein standard, but with anti-HNE antibody and HRP-labeled detection antibody. Use coating buffer instead of protein standard. 3. Negative controls for sandwich ELISA: (a) Blank-A: wells with capture antibody and known HNE-conjugated protein standard, but without the HRP-labeled anti-HNE antibody. Use the wash buffer B instead of the anti-HNE antibody. (b) Blank-B: for sandwich ELISA: wells with the capture antibody and HRP-labeled anti-HNE antibody, but without HNE-conjugated protein standard. Use PBS instead of the protein standard. 3.3 Indirect ELISA Method

1. Plan the layout of the plate and note the wells in which the HNE-conjugated protein standards, samples and controls will be added (see Note 10). 2. Prepare the sample and the optimized range of HNE-conjugated protein standards in the coating buffer (as previously determined through Subheading 3.1, steps 3 and 4). 3. Add 50 μL/well and incubate at RT for 2 h. 4. Aspirate the solution from the wells and wash wells with wash buffer B (see Note 11). 5. Add blocking buffer (300 μL/well) and incubate at RT for 1 h (see Note 12). 6. Wash wells with PBS. 7. Add the optimized concentration of anti-HNE antibody prepared in wash buffer A (50 μL/well). 8. Incubate at RT for 1 h. 9. Aspirate the solution from the wells and wash wells thrice with wash buffer B (see Note 11). 10. Prepare the optimized concentration of HRP-labeled detection antibody in wash buffer B and immediately wrap the tube in silver foil. 11. Add 50 μL of HRP labeled detection antibody per well, seal the plate with the plate sealer and immediately wrap it in a foil.

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12. Incubate at RT for 1 h in dark. 13. Aspirate the solution from the wells and wash wells thrice with wash buffer B and once with PBS (see Note 11). 14. Add TMB (100 μL/well) and incubate at RT for 20 min in dark. 15. Add 2 M sulfuric acid (100 μL/well) to stop the reaction (see Note 13). 16. Gently tap the plate to ensure thorough mixing. 17. Read the absorbance at 450 nm on a plate reader within 30 min of stopping the reaction. A positive sample develops a yellow color whereas a negative sample is colorless. 18. Compare the readings of the samples with the negative controls to confirm the presence of HNE in the sample and/or read off the concentration of HNE in the sample from the standard curve (see Note 14). 3.4 Sandwich ELISA Method

1. Plan the layout of the plate and note the wells in which the HNE-conjugated protein standards, samples and controls will be added (see Note 10). 2. Dilute the capture antibody in coating buffer and immediately add 100 μL/well. 3. Seal the plate with the plate sealer and incubate overnight at 4  C or at 37  C for 1 h 30 min. 4. Aspirate the solution from the wells and wash wells thrice with wash buffer A (see Note 11). 5. Add the blocking buffer (300 μL/well) and incubate at RT for 1 h (see Note 12). 6. Remove all blocking solution by inverting the plate in the sink and then wash wells gently with PBS. 7. Prepare the optimized range of HNE-conjugated protein standards in PBS (as determined previously through Subheading 3.1, step 3). 8. Add the standards and the optimized concentration of samples (as determined previously through Subheading 3.1, step 4) to the wells (50 μL/well). 9. Seal the plate with the plate sealer and incubate at 37  C for 2 h. 10. Aspirate the solution from the wells and wash wells twice with wash buffer A and once with PBS (see Note 11). 11. Prepare fresh HRP-labeled anti-HNE detection antibody in wash buffer B (at an optimized concentration as previously determined through Subheading 3.1, step 3) just before addition to wells and immediately wrap the tube in a foil (see Note 15).

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(a) Add the HRP-labeled detection antibody (50 μL/well). (b) Seal the plate, cover with foil and incubate 1 h at 37  C. (c) Aspirate the solution from the wells and wash wells thrice with wash buffer B and once with PBS (see Note 11). 12. Add TMB (100 μL/well) and incubate at RT for 20 min in dark. 13. Add 2 M sulfuric acid to stop the reaction (100 μL/well). (see Note 13) 14. Gently tap the plate to ensure thorough mixing. 15. Read the absorbance at 450 nm on a plate reader within 30 min of stopping the reaction. A positive sample develops yellow color whereas a negative sample is colorless. 16. Compare the readings of the samples with the negative controls to confirm the presence of HNE in the sample or read off the concentration of HNE in the sample from the standard curve (see Note 14).

4

Notes 1. Procurement can be carried out by a variety of companies. 2. If the recommended storage temperature for the antibodies is 20  C, then aliquot the stock in small aliquots to avoid repeated freeze–thaw cycles. These can be raised in a host species of choice such as rabbit, sheep or goat using HNE-KLH (keyhole limpet hemocyanin) as the antigen. The anti-HNE antibodies can be used as primary antibodies in indirect ELISA. 3. These are commercially available from several companies and should be against the host species in which the primary antibody (anti-HNE) is raised 4. If the detection antibody is conjugated to alkaline phosphatase, then p-nitrophenyl phosphate is used as a substrate and 3 M sodium hydroxide as a stop solution. The reaction is read at 405 nm. The volumes of the substrate and stop solution and the other procedures in the protocol remain unaltered. 5. Aliquot the stock solution of the HNE-protein standard in small volumes and store at 20  C. Avoid repeated freeze–thaw cycles. 6. Using special ELISA-grade plates is essential. Plates with curved bottoms allow efficient washing between steps and therefore reduced background. 7. To prepare the wash buffers, Tween should be added to PBS and allowed to mix on a shaker at RT for at least 1 h.

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8. Alternative blocking buffer can be used such as 1% BSA in 0.05% PBS-T (w/v). 9. Generally, detection of the antigen using the West PICO chemiluminescent kit recommends that the concentration of anti-HNE antibody is from 0.2 μg/mL to 1 μg/mL and the concentration of HRP-labeled detection antibody is from 0.01 μg/mL to 0.04 μg/mL. The secondary HRP-labeled detection antibody must be raised against the host species in which the anti-HNE antibody is raised. 10. To enable easy view of the wells during the addition of small volumes of antibody solutions, a colored piece of uniformly flat cardboard can be placed under the ELISA plate. 11. During all the washing steps, use 300 μL/well of the wash buffer (i.e., fill the wells up to the brim). During each washing step, the plate should be first inverted in the sink to remove most liquid. Then, the plate should be firmly blotted several times on a dry clean tissue paper until no liquid is visibly seen on the tissue. Complete removal of liquid from wells is essential before proceeding further to avoid poor reproducibility of results. 12. The volume of the blocking solution should be higher (at least double) than the volumes of the antibody solutions to allow complete and effective blocking. 13. The volume of stop solution should be equal to the volume of substrate solution. 14. Readings of the samples should be three times higher than the readings of the blank to obtain statistically significant differences. Determine the average absorbance readings for each set of triplicate standards, controls and samples. Normalize the readings to the blank i.e. subtract the reading of the blank (blank-2 and blank-B in indirect ELISA and sandwich ELISA, respectively) from that of the standards and samples. Construct a standard curve by plotting the concentration on the x-axis and the subtracted average absorbance for each standard on the y-axis. Draw a line of best fit through the points on the graph and determine the concentration of the unknown samples by using the equation of the line of regression. If diluted samples were used, then the concentration deduced from the standard curve should be multiplied by the dilution factor. Alternatively, a computer-based curve-fitting statistical software could be used to calculate the concentration of the sample. 15. In the sandwich ELISA format, if the anti-HNE antibody is not labeled with HRP, then an HRP-labeled antibody is required for detection. Thus, this format will involve three antibodies; capture, anti-HNE and the HRP-labeled detection antibody. The HRP-labeled antibody should be raised against the host

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species in which the anti-HNE antibody is raised and should bind exclusively to the anti-HNE antibody. It should be confirmed through indirect ELISA that this detection antibody does not bind to the capture antibody. For optimization of concentrations of the three antibodies, titrations should first be carried out with the capture and the anti-HNE antibody (Fig. 3), followed by optimization of the HRP-labeled detection antibody. References 1. Petersen DR, Doorn JA (2004) Reactions of 4-hydroxynonenal with proteins and cellular targets. Free Radic Biol Med 37:937–945 2. Lovell MA, Ehmann WD, Mattson MP, Markesbery WR (1997) Elevated 4-hydroxynonenal in ventricular fluid in Alzheimer’s disease. Neurobiol Aging 18:457–461 3. Kamimura S, Gaal K, Britton RS, Bacon BR, Triadafilopoulos G, Tsukamoto H (1992) Increased 4-hydroxynonenal levels in experimental alcoholic liver disease: association of lipid peroxidation with liver fibrogenesis. Hepatology 16:448–453 4. Shoeb M, Ansari NH, Srivastava SK, Ramana KV (2014) 4-hydroxynonenal in the pathogenesis and progression of human diseases. Curr Med Chem 21:230–237 5. Li CJ, Nanji AA, Siakotos AN, Lin RC (1997) Acetaldehyde-modified and 4-hydroxynonenal-

modified proteins in the livers of rats with alcoholic liver disease. Hepatology 26:650–657 6. Patel VB, Spencer CH, Young TA, Lively MO, Cunningham CC (2007) Effects of 4-hydroxynonenal on mitochondrial 3-hydroxy3-methylglutaryl (HMG-CoA) synthase. Free Radic Biol Med 43:1499–1507 7. Awasthi YC, Yang Y, Tiwari NK, Patrick B, Sharma A, Li J, Awashi S (2004) Regulation of 4-hydroxynonenal-mediated signaling by glutathione S-transferases. Free Radic Biol Med 37:607–619 8. Weber D, Milkovic L, Bennett SJ, Griffiths HR, Zarkovic N, Grune T (2013) Measurement of HNE-protein adducts in human plasma and serum by ELISA-comparison of two primary antibodies. Redox Biol 1:226–233

Chapter 5 Using Flow Cytometry to Detect and Measure Intracellular Thiol Redox Status in Viable T Cells from Heterogeneous Populations Alex J. Wadley, Rhys G. Morgan, Richard L. Darley, Paul S. Hole, and Steven J. Coles Abstract Increased production of reactive oxygen species (ROS) and deficiencies in cellular antioxidant defenses are the principal causes of cellular oxidative stress. ROS can react with a variety intracellular molecules, including redox active cysteine thiols (–SH) within proteins. Cysteine thiols can occupy several redox states and conversion between them is highly dynamic during, for example, cell growth, resulting in modification and subsequent loss of the “reduced thiol” form (–SH or –S
). The challenge lies with detecting and measuring thiol redox status inside viable heterogeneous cell populations (e.g., peripheral blood mononuclear cells (PBMCs)). Here we describe a flow cytometric approach for the evaluation of intracellular thiol redox status in human CD3+ T cells within a viable PBMC preparation. Using the thiol reactive probe, fluorescein-5 maleimide (F5M), we demonstrate that loss of reduced intracellular thiol correlates with a decrease in F5M fluorescence. We also detected a loss of F5M fluorescence in Jurkat cell cultures exposed to exogenous H2O2 generated by glucose oxidase. Since F5M binds irreversibly to reduced cysteine thiols, cells may be sorted based on F5M fluorescence intensity and redox active proteins can subsequently be extracted and separated using SDS-PAGE. This final step facilitates identification of redox active proteins from individual cell populations in live heterogeneous cell mixes using proteomic analysis. Key words Flow cytometry, Thiol, Redox, F5M, NEM, Glucose oxidase, Hydrogen peroxide, Oxidative stress, T cells, Jurkat

1

Introduction In mammals and plants, intracellular cysteine thiols play a key role in many cellular processes, including redox homeostasis [1] and signal transduction [2, 3]. Redox active solvent accessible cysteine residues have been identified in a variety of protein families including protein phosphatases (e.g., PTP1B [4]), transcription factors (e.g., Nf-κB [5]) and various metabolic enzymes (e.g., hBCATc [6]). In the case of hBCATc, thiol redox state can affect protein

John T. Hancock and Myra E. Conway (eds.), Redox-Mediated Signal Transduction: Methods and Protocols, Methods in Molecular Biology, vol. 1990, https://doi.org/10.1007/978-1-4939-9463-2_5, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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activity [7]. It is therefore plausible that alterations in the cellular redox environment could impact cell growth, function and survival [8]. Changes in the cellular redox environment can be mediated by the generation of reactive oxygen species (ROS). Indeed, certain ROS (e.g., H2O2) can modify cysteine thiols directly through oxidation [2]. The propensity of redox active proteins to become modified through cysteine oxidation will depend on the reduction mid-point potential of that protein, where alternate isoforms of the same protein can exhibit different redox sensitivities [9]. Reactive nitrogen species (RNS) are also known to modify cysteine thiols [10], increasing the complexity of the cellular system. Transient increases in ROS and RNS may occur during normal cell growth. Both ROS and RNS are unstable and have been shown undergo cross-reactions forming diverse unstable intermediates [11]. For example, H2O2 can form hydroxyl radicals (OH·), hypochloric acid (HOCl) and peroxynitrite (ONOO
) intermediates under physiological conditions via Fenton and Haber-Weiss reactions (reviewed in [12]). It is therefore challenging to interpret these data when examining the global effects of ROS and (or) RNS in viable cellular systems. Given the high potential for cross reactivity it is difficult to determine which particular species are targeting the cysteine thiols in cells. In such systems it may be more appropriate for the researcher to interpret their findings with respect to the availability of “reduced” cysteine thiols. A further complexity is observed when heterogeneous cell populations such as peripheral blood mononuclear cells (PBMC) are used for in vitro/ex vivo models. In this scenario, the researcher may wish to evaluate these effects for a particular cell type (e.g., T cells) within the viable PBMC mix whilst retaining the integrity of the in vitro model. To overcome these technical difficulties we describe a flow cytometric method which quantitates the availability of reduced cysteine thiols using a fluorescent probe. We show that this method can be used to measure the intracellular availability of reduced cysteine thiols for viable T cells that exist within a PBMC population. Flow cytometric methods that incorporate methyl orange for the analysis of low molecular weight thiols, such as glutathione, have been described [13]. However, our method incorporates the thiol reactive probe fluorescein-5 maleimide (F5M), as shown in Fig. 1, which will also label redox active proteins with reduced solvent accessible cysteines [14]. Fluorescein derivatives will freely enter cells with maximum accumulation achieved within 20–30 min [15]. As such, compounds such as fluorescein diacetate are commonly used in flow cytometric viability staining panels and are detected in the FL1 parameter (laser: λ488 nm, filter: λ530/30 nm) [16]. In the method described here, F5M accumulation in cells is confirmed using confocal microscopy and nonspecific F5M mediated fluorescence is determined using N-ethylmaleimide pretreatment, which masks all solvent accessible reduced thiols

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Fig. 1 Molecular structures of N-ethylmaleimide (NEM) and fluorescein-5 maleimide (F5M). The maleimide alkene (C–C double bond) participates in a Michael addition with strong nucleophiles such as cysteine thiol (or thiolate) groups (–SH or –SO
), creating an irreversible thioether bond (C–S). On comparing NEM with F5M, a substitution of “ethane” with “fluorescein” at the functional imide group is noted

(Fig. 1). The flow cytometric strategy described here is extremely powerful; since F5M only occupies one detection parameter (FL1), the remaining fluorescent detection parameters are available for cell phenotype and functional analysis. To demonstrate this power we show that T cells can be identified in the FL4 parameter (laser: λ640 nm, filter: λ675/25 nm) in conjunction with F5M analysis. Furthermore, the method may also be used to monitor intracellular oxidative stress mediated by exogenous H2O2 in viable cells, as we demonstrate using an in vitro model. For a final step, cells labeled with F5M can be sorted and redox active proteins identified by SDS-PAGE. The flexibility of this technique allows study of redox changes within various heterogeneous viable cell populations, (e.g., those that occur in PBMCs during periods of intense exercise) [17]. This method may also be clinically important for diseases such as acute myeloid leukemia where ROS and intracellular redox status are implicated [18, 19]. The method presented here has been adapted or otherwise reproduced from our previous work “Detecting intracellular thiol redox state in leukaemia and heterogeneous immune cell populations: An optimised protocol for digital flow cytometers” by Wadley et al., published under CC BY 4.0 [20].

2

Materials 1. Complete IMDM tissue culture medium: containing 10% (v/v) fetal calf serum (FCS), 2 mM L-glutamine, 100 U/mL penicillin, and 100 mg/mL streptomycin.

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2. Hanks’ balanced salt solution. 3. Dulbecco’s phosphate buffered saline (D-PBS) endotoxin tested, pH 7.0 (Sigma-Aldrich). 4. Flow stain buffer (FSB): D-PBS adjusted to contain 2% (v/v) FCS, 0.02% NaN3 (v/v) and 5 mM EDTA (stored at 4  C). 5. Ficoll-Paque PLUS. 6. Trypan blue solution 0.4%. 7. Jurkat cells (LCG Standards). 8. Mouse anti-human CD3 (clone SK7) APC conjugate. 9. Mouse IgG1 APC conjugate isotype control. 10. Glucose oxidase from Aspergillus niger: 100,000 U/g. 11. Catalase. 12. N-Ethylmaleimide (NEM): 1 mM stock in D-PBS stored at 4  C protected from light. 13. Fluorescein-5 maleimide (F5M): 1 mM stock in D-PBS stored at 4  C protected from light. 14. Cell extraction buffer: 10 mL stock of 50 mM Tris, pH 7.4 adjusted to contain, 0.8 M NaCl, 5 mM MgCl2, 1 mM EDTA, 0.5% (v/v) NP-40 and 1 complete mini-EDTA free protease inhibitor tablet, filtered and stored at 4  C. 15. Novex® 4–20% gradient precast tris-glycine gel. 16. NuPAGE® MOPS SDS-PAGE running buffer. 17. NuPAGE® LDS SDS-PAGE sample loading buffer. 18. G-250 colloidal coomassie blue reagent. 19. SDS-PAGE gel fix solution: 50% (v/v) methanol, 10% (v/v) acetic acid. 20. Multiparameter flow cytometer.

3

Methods

3.1 Separation of Peripheral Blood Mononuclear Cells from Whole Venous Blood

Venous blood from healthy donors is collected by a trained phlebotomist into a 6 mL lavender stopper K2EDTA Vacationer® tube. All procedures from this point on are performed inside a Class 2 biological safety cabinet. It is important that the Class 2 biological safety cabinet in switched on, swabbed with 70% ethanol and allowed to circulate for at least 30 min prior to any venous blood work (see Note 1). 1. Transfer 6 mL venous blood to a fresh sterile universal tube and add an equal volume of Hanks’ balanced salt solution (HBS); 12 mL total volume (see Note 2).

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2. Add 3 mL Ficoll-Paque PLUS to three sterile 15 mL centrifuge tubes and carefully layer 4 mL of the diluted venous blood on top of the Ficoll-Paque PLUS (see Note 3). 3. Transfer the tubes to a bench-top centrifuge with swing-out rotor and centrifuge at 400  g for 40 min at room temperature (18–20  C) with the brake off (see Note 4). 4. Carefully remove the tubes from the centrifuge and return to the biological safety cabinet. The PBMC layer (white and delicate) will be visible at the Ficoll-Paque PLUS interface. Carefully remove the upper layer (plasma) to waste with a fresh sterile Pasteur pipette without disturbing the PBMC layer. Then using a fresh sterile Pasteur pipette, transfer the PBMC layer to a fresh 15 mL centrifuge tube. For each tube, remove the entire PBMC interface with as little of the FicollPaque PLUS layer as possible. 5. Add 8 mL HBS to each tube and centrifuge at 60–100  g for 10 min at room temperature. 6. Using a fresh sterile pipette, remove the supernatant from each tube taking great care not to disturb the PBMC pellet and suspend in 1 mL HBS. The PBMCs can be pooled at this stage. Add sufficient HBS to the pooled cells to make 10 mL. 7. Centrifuge pooled PBMCs at 60–100  g for 10 min at room temperature. 8. Remove the supernatant taking great care not to disturb the PBMC pellet and suspend the pellet in 2 mL prewarmed (37  C) complete IMDM. Transfer the cells to an incubator set at 37  C and 5% CO2 for subsequent analysis (see Note 5). 3.2 Flow Cytometric Analysis of T Cell Intracellular Reduced Thiol Within the PBMC Preparation

Following preparation of PBMC as described above, intracellular reduced thiol analysis in viable T cells can now be performed. Prepare the F5M working solution for flow cytometric use in advance. Protect from light and store at 4  C (up to 1 month) or at
20  C for up to 1 year (see Note 6). 1. Remove the PBMCs from the CO2 incubator and centrifuge at 60–100  g in a benchtop refrigerated centrifuge with swingout rotor for 5 min at room temperature. 2. Remove supernatant and suspend the pellet in 10 mL D-PBS (equilibrated to room temperature) and centrifuge at 60–100  g for 10 min. 3. Remove supernatant and suspend the pellet in 1 mL FSB. Prepare a 1 in 20 dilution of the PBMC suspension for cell viability counting (5 μL PBMC in 95 μL FSB is sufficient) and transfer the remaining suspension to ice (see Note 7). 4. Mix 10 μL of the 1 in 20 PBMC dilution with 10 μL trypan blue solution and count viable cells using a hemocytometer (see Note 8).

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5. Adjust the PBMC suspension (on ice from step 3) to a density of 107 viable cells/mL with prechilled (4  C) FSB and add 100 μL (106 cells) to three fresh 15 mL centrifuge tubes and centrifuge for 5 min at 60–100  g and 4  C (see Note 9). 6. Remove supernatants and suspend each pellet in FSB to a final volume of 90 μL. 7. Add 10 μL of mouse anti-human CD3 (clone SK7) APC conjugate to two tubes and 10 μL of mouse IgG1 APC conjugate isotype control to the other. 8. Vortex all tubes briefly (1–2 s) and incubate on ice for 30 min protected from the light. 9. Add 1 mL D-PBS (chilled to 4  C) to each tube and centrifuge for 5 min at 60–100  g and 4  C. 10. Remove supernatants and suspend each pellet in 1 mL D-PBS and centrifuge for a further 5 min at 60–100  g and 4  C. 11. Remove supernatants and add 1 mL D-PBS to the isotype control tube and 1 mL D-PBS to one of the anti-human CD3 tubes and label it “F5M.” 12. Add 1 mL of 1 μM NEM solution to the remaining anti-human CD3 tube. These cells will be used to set the background for the F5M T cell stain. Label this tube “NEM.” 13. Incubate all tubes for 20 min on ice, protected from the light. 14. Centrifuge for 5 min at 60–100  g and 4  C. 15. Remove supernatants and suspend each pellet in 1 mL D-PBS and centrifuge for 5 min at 60–100  g and 4  C. 16. Remove supernatants and add 1 mL chilled D-PBS to the isotype control tube. These cells will be used to set the negative T cell gate. Add 1 mL 0.1 μM F5M to both “NEM” and “F5M” tubes. 17. Incubate all three tubes for 20 min on ice, protected from the light. 18. Centrifuge for 5 min at 60–100  g and 4  C. 19. Remove supernatants and suspend each pellet in 1 mL prechilled FSB and centrifuge for 5 min at 60–100  g and 4  C. Repeat this step. 20. Finally remove supernatants and suspend each pellet in 100 μL prechilled FSB, keep on ice, protect from light and go to flow cytometer. 21. At the flow cytometer, acquire the “Ig-control” cells first. These cells can be used to set the negative T cell gate, which will appear in the FL4 parameter (laser: λ640 nm, filter: λ675/25 nm). Figure 2a, b illustrates the gating strategy.

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22. Next acquire the “NEM” cells and create an FL1 (laser: λ488 nm, filter: λ530/30 nm) histogram plot gated on the T cell population as seen in Fig. 2b. The peak shown on the FL1 histogram is representative of the F5M background staining and can be seen in Fig. 2c (see Note 10). 23. Finally acquire the “F5M” cells. As shown in Fig. 2c, these cells fluoresce at an order of magnitude greater than “NEM” cells. 24. To confirm viability, perform cell counts after flow cytometry acquisition as described in steps 3 and 4 (see Note 11). Figure 2d shows little loss in cell viability post treatment. 25. As shown in Fig. 2e, f, confocal microscopy can be used to confirm that the F5M is entering the T cells and therefore that the flow cytometric FL1 signal generated is representative of intracellular thiol redox status (see Note 12). 3.3 Analyzing the Effect of Extracellular H2O2 on Intracellular Thiol Oxidation Using Flow Cytometry

The method described in Subheading 3.2 can be taken further and used to monitor intracellular thiol redox status in viable cells that are exposed to exogenous ROS. This technique is of particular importance, since some ROS (e.g., H2O2) have the capacity to cross cell membranes (reviewed in [21]). This exogenous H2O2 then has potential to enter neighboring cells, including T cells which are central mediators in disease immunity (reviewed in [22, 23]). Exposure to a persistent source of exogenous H2O2 can be modeled in vitro by adding GOX to the culture medium [19]. The Jurkat immortalized T cell line was used to demonstrate this principle for this section. As outlined in Subheading 3.1, a robust aseptic technique must be observed throughout this procedure. 1. Maintain Jurkat cells in 15 mL complete IMDM, at a density between 2  105 and 106 cells/mL at 37  C and 5% CO2 (see Note 13). 2. Prepare a 10 mU/mL GOX solution and a 10 mg/mL catalase solution and store at 4  C until required (see Note 14). 3. Transfer the contents of a 24–48 h Jurkat cell culture flask to a 15 mL centrifuge tube and centrifuge at 60–100  g for 10 min at room temperature in a bench-top centrifuge with swing-out rotor. 4. Remove the supernatant and carefully suspend the pellet in 1 mL complete IMDM prewarmed to 37  C (see Note 15). 5. Perform a viable cell count as described in Subheading 3.2, steps 5–7 and dilute the Jurkat cells to a viable density of 107 cells/mL. 6. Add 100 μL of the diluted Jurkat cell suspension to five wells in a 96-well tissue culture treated plate, plus 98 μL to two wells in the same plate and label; “0 h,” “1 h,” “2 h,” “3 h,” “4 h,” “0 h +catalase,” and “4 h +catalase” accordingly (see Note 16).

Fig. 2 Flow cytometric analysis of intracellular thiol redox status in primary human T cells. PBMCs were prepared from venous blood using Ficoll-Paque density centrifugation and washed in D-PBS. Washed PBMCs (106 cells in 100 μL) were labeled with anti-human CD3-APC. PBMCs were then treated with either 1 μM NEM followed by 0.1 μM F5M, or 0.1 μM F5M alone and analyzed by flow cytometry. (a) Representative forward light scatter (FSC) vs. side light scatter (SSC) flow cytometric profile with the treated PBMCs identified in P2. (b) Gated on P2, CD3+ T cells are shown in the FL4 parameter (laser: λ640 nm, filter: λ675/25 nm) in R1 (c). Panel shows representative fluorescence histograms depicting F5M fluorescence in control and treated T cells (gated on R1). (d) T cell viability was monitored before and after NEM and F5M treatment using trypan blue staining. Confocal microscopy was used to confirm (e) F5M cellular uptake in PBMCs and (f) surface staining for T cells with anti-human CD3-APC. Figure taken from “Detecting intracellular thiol redox state in leukaemia and heterogeneous immune cell populations: An optimised protocol for digital flow cytometers” by Wadley et al., used under CC BY 4.0 [20]

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Label the wells that contain 98 μL “0 h +catalase” and “4 h +catalase” to account for the addition of catalase. 7. Add 2 μL of the 10 mg/mL catalase stock solution (prepared in step 2) to wells labeled “0 h +catalase” and “4 h +catalase” only (see Note 17). 8. To start the assay add 100 μL of the GOX solution (prepared in step 2) to wells labeled “4 h” and “4 h +catalase” and incubate at 37  C and 5% CO2. 9. After 1 h add 100 μL of the GOX solution to the “3 h” well and return to the incubator. 10. Repeat step 10 after a further 1 and 2 h for wells labeled “2 h” and “1 h” respectively. 11. Finally, 1 h after the addition of GOX to “1 h,” add 100 μL of the GOX solution to wells labeled “0 h” and “0 h +catalase.” Immediately transfer the contents of each well to a fresh 1.5 mL microcentrifuge tube and centrifuge all at 60–100  g for 5 min at room temperature. 12. Carefully remove supernatants and suspend the pellets in 1 mL D-PBS. Centrifuge all tubes at 60–100  g for 5 min at room temperature. 13. Remove supernatants and suspend pellets in 1 mL prechilled 0.1 μM F5M solution and incubate all tubes for 20 min, on ice and protected from light. 14. Centrifuge tubes at 60–100  g for 5 min at 4  C. 15. Remove supernatants and suspend pellets in 1 mL prechilled FSB and centrifuge at 60–100  g for 5 min at 4  C. 16. Remove supernatants and suspend pellets in 100 μL FSB and go to flow cytometer. Keep tubes on ice and protect from light. 17. At the flow cytometer, acquire the “0 h” tube first, followed by the others in chronological order. Draw a region (P2) around the Jurkat cell population as seen in Fig. 3a and create an FL1 histogram gated on the P2 region as seen in Fig. 3b. 18. Normalize the FL1 mean fluorescent intensity (MFI) values to “0 h” in order interpret the flow cytometry data as shown in Fig. 3c and in Table 1. 19. Viable cell counts can be performed on GOX treated and F5M labeled cells prior or following flow cytometry as shown in Fig. 4a. 20. To demonstrate that the GOX generated H2O2 is mediating the loss in reduced thiols, the effect of catalase in “4 h” vs. “4 h +catalase” treated cells can be observed on normalized data as shown in Fig. 4b. The data suggest that H2O2 mediates the loss in reduced intracellular thiols as measured by a loss in F5M (FL1) fluorescent signal.

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A

B 0h 3h

2h

4h 1h

Intracelluar thiol redox state (normalised to 0h)

C

2 1.5 Reduced

1

Oxidised 0.5 0

0

1

2 3 GOX incubation time (h)

4

Fig. 3 The effect of exogenous GOX-generated H2O2 on the loss of intracellular reduced thiol using an in vitro immortalized Jurkat T cell model. (a) Representative FSC vs SSC flow cytometric profile of GOX treated Jurkat cells (P2). (b) Histogram (gated on P2) illustrates loss of F5M signal in Jurkat cells following incubation with 5 mU/mL GOX for 0–4 h. Samples were taken for flow cytometric analysis after the following times; 0 h, 1 h, 2 h, 3 h and 4 h. Data were compared with 0 h to normalize the loss of F5M signal (see Table 1). (c) Chart illustrates normalized FL1 (laser: λ488 nm, filter: λ530/30 nm) MFI. An increase in F5M signal is noted at 1 h (suggesting a reductive spike), followed by progressive decrease in signal at 2–4 h (suggesting a loss of reduced thiols). Data shown are mean  SD of the mean (n ¼ 3). Figure taken from “Detecting intracellular thiol redox state in leukaemia and heterogeneous immune cell populations: An optimised protocol for digital flow cytometers” by Wadley et al., used under CC BY 4.0 [20] 3.4 Analyzing F5M Labeled Proteins Extracted from Viable Cells

Treating viable cells with F5M will result in the labeling of solvent accessible intracellular “reduced” cysteine thiols [14]. This will include low molecular weight thiols such as glutathione [24], however some of these reduced cysteine thiols will be present in the native conformation of redox active proteins. It would be expected that if a reduced cysteine thiol is lost through oxidation, F5M labeling will not occur. This will result in a decreased fluorescent signal relating to a particular redox active protein when separated using SDS-PAGE. Cells treated in Subheading 3.2 or 3.3 can

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Table 1 Raw flow cytometry data and normalisednormalized thiol redox state. Table taken from “Detecting intracellular thiol redox state in leukaemia and heterogeneous immune cell populations: An optimised protocol for digital flow cytometers” by Wadley et al., used under CC BY 4.0 [20] GOX incubation time (h)

Mean fluorescence intensity (MFI) in FL1 (arbitrary units)

Normalized intracellular thiol redox state (later time/0 h)a

0

3,163,900.56

0 h MFI/0 h MFI ¼ 1

1

5,409,960.52

1 h MFI/0 h MFI ¼ 1.64 (p