Nitric Oxide: Methods and Protocols [1 ed.] 1617379638, 9781617379635

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ME T H O D S

IN

MO L E C U L A R BI O L O G Y

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

For other titles published in this series, go to www.springer.com/series/7651

TM

Nitric Oxide Methods and Protocols

Edited by

Helen O. McCarthy School of Pharmacy, Queen’s University Belfast, Northern Ireland, UK

Jonathan A. Coulter School of Pharmacy, Queen’s University Belfast, Northern Ireland, UK

Editors Helen O. McCarthy School of Pharmacy Queen’s University, Belfast Northern Ireland BT9 7BL, UK [email protected]

Jonathan A. Coulter School of Pharmacy Queen’s University, Belfast Northern Ireland BT9 7BL, UK [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-61737-963-5 e-ISBN 978-1-61737-964-2 DOI 10.1007/978-1-61737-964-2 Springer New York Dordrecht Heidelberg London © Springer Science+Business Media, LLC 2011 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface This book forms part of the highly acclaimed Methods in Molecular BiologyTM series, which aims to provide a detailed reference manual giving a step-by-step approach to reproduce various complex protocols within your own laboratory. For each volume in this series, editors have included the most interesting and relevant methodologies published in the field in recent years, thereby providing access to the most novel experimental approaches. In addition, this series also provides a detailed notes section which documents specific information relating to particularly challenging aspects of a methodology. The past two decades have seen an explosion in the number of research articles relating to both the physiological and pathological responses evoked by nitric oxide generation. Despite this, accurate quantification of nitric oxide in either in vitro or in vivo models remains challenging, due to the relatively unstable nature of the molecule. This volume considers two of the main aspects of nitric oxide research. Section I of the book includes a review from an expert in tumor radiosensitization induced by various novel compounds, including nitric oxide. The review covers multiple facets of nitric oxide including its role in addiction, the cardiovascular system, the nervous system, and cancer. The remainder of Section I describes various disparate protocols relating to the direct detection and quantification of nitric oxide. These include techniques which detail how to image real-time in vivo generation of nitric oxide, quantify nitric oxide production in the rat brain, and detect ultralow levels of nitric oxide in the pM range. Section II focuses primarily on techniques designed to either inhibit or enhance nitric oxide, with an aim to achieve therapeutic gain. These include inhibition of the nitric oxide synthase enzymes using viral, shRNA delivery systems to prevent cardiovascular dysfunction, peripheral neuropathy, and graft rejection. Other techniques highlighted deal with the overproduction of nitric oxide at target sites using novel nitric oxide releasing nanoparticles and biofilms. We hope this book provides clarification on the numerous complex methodologies detailed in each chapter, proving to be an invaluable resource for anyone with an interest in nitric oxide research. Helen O. McCarthy Jonathan A. Coulter

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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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. . . . . . . . . . . . . . . . . . . . . .

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Nitric Oxide Physiology and Pathology David G. Hirst and Tracy Robson

SECTION I 2.

3.

4.

5.

6.

DETECTION AND QUANTIFICATION OF NITRIC OXIDE

Ionizing Radiation-Induced DNA Strand Breaks and γ-H2AX Foci in Cells Exposed to Nitric Oxide . . . . . . . . . . . . . . . . . . . . . . . . . . . Kai Rothkamm and Susanne Burdak-Rothkamm

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Determination of S-Nitrosothiols in Biological Fluids by Chemiluminescence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Enika Nagababu and Joseph M. Rifkind

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Measurement of Nitrite in Blood Samples Using the Ferricyanide-Based Hemoglobin Oxidation Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . Barbora Piknova and Alan N. Schechter

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Selective Fluorescent Activation for Bioimaging the Expression of Nitric Oxide in Cellular and In Vivo Systems . . . . . . . . . . . . . . . . . . . . . . . Junfeng Zhang and Hao Hong

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Real-Time Measurement of Murine Hippocampus NO Levels in Response to Cerebral Ischemia/Reperfusion . . . . . . . . . . . . . . . . . . . . . . . . . Xiaoxiang Zheng, Kezhou Liu, and Yong Yang

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Detection of Low Levels of Nitric Oxide Using an Electrochemical Sensor . . . . Yong Chool Boo, Gyeong In Mun, Sarah L. Tressel, and Hanjoong Jo

8.

Determination of the Scavenging Capacity Against Reactive Nitrogen Species by Automatic Flow Injection-Based Methodologies . . . . . . . . . . . . Marcela A. Segundo, Luís M. Magalhães, Joana P.N. Ribeiro, Marlene Lúcio, and Salette Reis

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Aqueous Measurement of Nitric Oxide Using Membrane Inlet Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 David N. Silverman and Chingkuang Tu

10. Quantum Cascade Laser Technology for the Ultrasensitive Detection of Low-Level Nitric Oxide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 Angela Elia, Pietro Mario Lugarà, Cinzia Di Franco, and Vincenzo Spagnolo

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11. Determination of In Vivo Nitric Oxide Levels in Animal Tissues Using a Novel Spin Trapping Technology . . . . . . . . . . . . . . . . . . . . . . . . . 135 Anatoly F. Vanin and Alexander A. Timoshin SECTION II

NITRIC OXIDE GENERATION

12. β Cell Protection by Inhibition of iNOS Through Lentiviral Vector-Based Strategies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153 Sean O. Hynes, Cillian McCabe, and Timothy O’Brien 13. Characterization of Nitric Oxide Delivery Systems Produced By Various Nanotechnologies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169 Chi H. Lee 14. Nitric Oxide Releasing Nanoparticle Synthesis and Characterization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187 George Han, Adam J. Friedman, and Joel M. Friedman 15. NOS Antagonism Using Viral Vectors as an Experimental Strategy: Implications for In Vivo Studies of Cardiovascular Control and Peripheral Neuropathies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 197 Beihui Liu, James Hewinson, Haibo Xu, Francisco Montero, Carmen R. Sunico, Federico Portillo, Julian F.R. Paton, Bernardo Moreno-López, and Sergey Kasparov Index

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 225

Contributors YONG CHOOL BOO • Department of Molecular Medicine and Cell and Matrix Research Institute, BK21 Medical Education Program for Human Resources, Kyungpook National University School of Medicine, Daegu, Republic of Korea SUSANNE BURDAK-ROTHKAMM • Stoke Mandeville Hospital, Histopathology Department, Aylesbury, UK CINZIA DI FRANCO • CNR-IFN U.O.S. di BARI, Physics Department, University of Bari, Bari, Italy ANGELA ELIA • CNR-IFN U.O.S. di BARI, Physics Department, University of Bari, I-0126 Bari, Italy JOEL M. FRIEDMAN • Department of Physiology and Biophysics, Albert Einstein College of Medicine, Yeshiva University, Bronx, NY ADAM J. FRIEDMAN • Division of Dermatology, Department of Physiology and Biophysics, Albert Einstein College of Medicine, Yeshiva University, Bronx, NY GEORGE HAN • Department of Physiology and Biophysics, Albert Einstein College of Medicine, Yeshiva University, Bronx, NY JAMES HEWINSON • Department of Physiology and Pharmacology, University of Bristol, Bristol, UK DAVID G. HIRST • School of Pharmacy, Queen’s University Belfast, Belfast, UK HAO HONG • The State Key Laboratory of Pharmaceutical Biotechnology, School of Life Sciences, Nanjing University, Nanjing, People’s Republic of China SEAN O. HYNES • Regenerative Medicine Institute, National University of Ireland, Galway, Ireland HANJOONG JO • Wallace H. Coulter Department of Biomedical Engineering at Georgia Tech, Emory University and Ewha Womans University, Atlanta, GA, USA SERGEY KASPAROV • Department of Physiology and Pharmacology, University of Bristol, Bristol, UK CHI H. LEE • School of Pharmacy, University of Missouri, Kansas City, MO, USA BEIHUI LIU • Department of Physiology and Pharmacology, University of Bristol, Bristol, UK KEZHOU LIU • Department of Biomedical Engineering, Zhejiang University, Hangzhou, People’s Republic of China MARLENE LÚCIO • REQUIMTE, Departamento de Química-Física, Faculdade de Farmácia, Universidade do Porto, Porto, Portugal PIETRO MARIO LUGARÀ • CNR-IFN U.O.S. di BARI, Physics Department, University of Bari, Bari, Italy LUÍS M. MAGALHÃES • REQUIMTE, Departamento de Química-Física, Faculdade de Farmácia, Universidade do Porto, Porto, Portugal CILLIAN MCCABE • Regenerative Medicine Institute, National University of Ireland, Galway, Ireland FRANCISCO MONTERO • Área de Fisiología, Facultad de Medicina, Universidad de Cádiz, Cádiz, Spain

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BERNARDO MORENO-LÓPEZ • Área de Fisiología, Facultad de Medicina, Universidad de Cádiz, Cádiz, Spain GYEONG IN MUN • Department of Molecular Medicine, Cell and Matrix Research Institute, BK21 Medical Education Program for Human Resources, Kyungpook National University School of Medicine, Daegu, Republic of Korea ENIKA NAGABABU • Molecular Dynamics Section, National Institute on Aging, National Institutes of Health, Baltimore, MD, USA TIMOTHY O’BRIEN • Regenerative Medicine Institute, National University of Ireland, Galway, Ireland JULIAN F.R. PATON • Department of Physiology and Pharmacology, University of Bristol, Bristol, UK BARBORA PIKNOVA • Molecular Medicine Branch, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, MD, UK FEDERICO PORTILLO • Área de Fisiología, Facultad de Medicina, Universidad de Cádiz, Cádiz, Spain SALETTE REIS • REQUIMTE, Departamento de Química-Física, Faculdade de Farmácia, Universidade do Porto, Porto, Portugal JOANA P.N. RIBEIRO • REQUIMTE, Departamento de Química-Física, Faculdade de Farmácia, Universidade do Porto, Porto, Portugal JOSEPH M. RIFKIND • Molecular Dynamics Section, National Institute on Aging, National Institutes of Health, Baltimore, MD, USA TRACY ROBSON • School of Pharmacy, Queen’s University Belfast, Belfast, UK KAI ROTHKAMM • Health Protection Agency Centre for Radiation, Chemical & Environmental Hazards, Oxon, UK; Stoke Mandeville Hospital, Histopathology Department, Aylesbury, UK; Radiation Protection Division, Health Protection Agency, Chilton, Didcot, UK ALAN N. SCHECHTER • Molecular Medicine Branch, National Institute of Diabetes, Digestive, Kidney Diseases, National Institutes of Health, Bethesda, MD, UK MARCELA A. SEGUNDO • REQUIMTE, Departamento de Química-Física, Faculdade de Farmácia, Universidade do Porto, Porto, Portugal DAVID N. SILVERMAN • Department of Pharmacology, University of Florida, Gainesville, FL VINCENZO SPAGNOLO • CNR-IFN U.O.S. di BARI, Physics Department, University of Bari, Bari, Italy CARMEN R. SUNICO • Área de Fisiología, Facultad de Medicina, Universidad de Cádiz, Cádiz, Spain ALEXANDER A. TIMOSHIN • Institute of Experimental Cardiology, Russian Cardiology Research-and-Production Complex, Rosmedtechnology Corporation, Moscow, Russia SARAH L. TRESSEL • Wallace H. Coulter Department of Biomedical Engineering at Georgia Tech, Emory University, Atlanta, GA, USA CHINGKUANG TU • Department of Pharmacology, University of Florida, Gainesville, FL ANATOLY F. VANIN • Semyonov Institute of Chemical Physics, Russian Academy of Sciences, Moscow, Russia HAIBO XU • Department of Pharmacology, State Key Laboratory for Research and Development of Chinese Materia Medica, Chengdu University of Traditional Chinese Medicine, Chengdu, P.R. China

Contributors

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YONG YANG • Department of Biomedical Engineering, Hangzhou Dianzi University, Hangzhou, People’s Republic of China JUNFENG ZHANG • The State Key Laboratory of Pharmaceutical Biotechnology, School of Life Sciences, Nanjing University, Nanjing, People’s Republic of China XIAOXIANG ZHENG • Department of Biomedical Engineering, Zhejiang University, Hangzhou, People’s Republic of China

Section I Detection and Quantification of Nitric Oxide

Chapter 1 Nitric Oxide Physiology and Pathology David G. Hirst and Tracy Robson Abstract Nitric oxide (NO) is just one member of a new class of gaseous signalling molecules with fundamental actions in biology. In higher vertebrates it has key roles in maintaining haemostasis and in smooth muscle (especially vascular smooth muscle), neurons and the gastrointestinal tract. It is intimately involved in regulating all aspects of our lives from waking, digestion, sexual function, perception of pain and pleasure, memory recall and sleeping. Finally, the way it continues to function in our bodies will influence how we degenerate with age. It will likely play a role in our deaths through cardiovascular disease, stroke, diabetes and cancer. Our ability to control NO signalling and to use NO effectively in therapy must therefore have a major bearing on the future quality and duration of human life. Key words: Nitric oxide, cardiovascular, cancer, wound healing.

1. Only One of the Gaseous Messengers

It was long believed that signalling within and between cells in the body is mediated by complex molecules such as proteins and peptides. It is becoming increasingly clear, however, that another class, the small gaseous messengers, plays crucial signalling roles in most, if not all, tissues. These molecules have been termed “gasotransmitters” (1) and include nitric oxide (NO), carbon monoxide (CO) and hydrogen sulphide (H2 S). Specific targets have been identified in smooth muscle cells, neurons and the gastrointestinal tract (2). It is likely that other molecules in this class will be identified; however, while new pathways await discovery for all of these gaseous molecules, NO is by far the most thoroughly studied and a truly enormous body of information has been

H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_1, © Springer Science+Business Media, LLC 2011

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accumulated (over 100,000 citations in the biomedical literature alone). On initial consideration, a more unlikely candidate for the regulation of normal physiology is hard to imagine. Until the 1980s, the free radical NO was noteworthy only for its influence as a constituent of smog and its only physiological role was as a respiratory irritant (3). All that changed with the observation that the “respiratory burst” by which macrophages kill pathogenic bacteria and cancer cells is arginine dependent (4). Soon after, NO was specifically identified as one of the most important cytotoxic molecules released by phagocytic leucocytes (5). In this setting, the reactivity of NO as a free radical made sense and this cytotoxic property has led to the development of NO generating systems for use in cancer therapeutics (6). However, around the same time other roles for controlled release of NO were identified and we now know that NO performs key regulatory functions in most tissues.

2. Regulation of Respiration There is evidence that NO concentrations in the nanomolar range, as generated by the constitutively expressed isoform of nitric oxide synthase (eNOS), can inhibit the affinity of cytochrome C oxidase for oxygen and so regulate cellular respiration rate (7). Inhibition of the respiratory chain by high NO concentrations resulting from activity of the inducible isoform of the enzyme (iNOS) may be permanent, because it leads to generation of highly damaging nitrosative stress (8). These mechanisms are complex, because even at high NO concentrations cytochrome C oxidase is protected against complete inactivation by NO (9). NO has also been shown to promote the formation of mitochondria and so enhance the cell’s capacity for oxidative metabolism (10). These processes have been implicated in a variety of pathologies including neurodegeneration (11).

3. Cardiovascular Effects NO is a key functional regulator in the cardiovascular system with roles extending beyond its initially identified action as a vasodilator: it controls vascular smooth muscle cell proliferation and migration, fibrinolysis, the adhesion of platelets and white blood cells and angiogenesis.

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3.1. Vascular Smooth Muscle Tone

The importance of NO in the vasculature was first realized when it was revealed to be indistinguishable from the endotheliumderived relaxing factor (EDRF) released when blood vessels are exposed to acetylcholine (12–15). A common feature of many of NO’s functions is that they are regulated by the second messenger guanylate cyclase, which in turn activates cyclic GMP. The physiological responses are then mediated by the interaction with cGMP-dependent kinases. This pathway requires only picomolar– nanomolar concentrations of NO which can be effectively generated by the constitutively expressed isoforms of NOS. This reaction requires as a substrate the amino acid arginine together with cofactors including tetrahydrobiopterin (BH4 ), NADH and molecular oxygen. Activation of this mechanism in blood vessels occurs either through the release of acetylcholine at parasympathetic nerve endings or signalling activated by shear stress recognition by the endothelium, both of which result in an increase in calcium flux. While there is little doubt that NO generated from eNOS is the main endothelium-derived relaxing factor in large vessels as originally proposed (13–15), there is evidence that this mechanism is not the sole NO generating pathway and that another NOS isoform, nNOS, can also catalyze the reaction in large vessels (16, 17). nNOS has also been shown to have distinct roles in the human coronary vascular bed (18). Even the classical model whereby the endothelium, in response to neurotransmitters or physical forces, releases NO may not be the dominant mechanism in all vascular beds. A recent study of mesenteric arteries in the toad showed that NO-mediated dilatation originated from nitrergic nerves rather than the endothelium (19). This is consistent with a previous observation that nNOS mediates cerebral vasodilation (20). Different mechanisms again probably exist in the microvasculature. It is well established that binding to the heme moiety of haemoglobin is a principal mechanism limiting the lifetime of NO in tissue (21), but it has also been proposed that haemoglobin may act as a reservoir for NO and its release can be stimulated by low oxygen tensions that induce allosteric changes in the haemoglobin molecule (22). It has also been shown that under hypoxic conditions reactions involving the reduction of nitrite by xanthine oxidase (23) or aldehyde oxidase (24) can lead to significant NO generation.

3.2. Cell Adhesion to the Endothelium

Adhesion of leucocytes to blood vessel walls is an important component of the response that targets inflammatory cells to sites of infection; however, an inappropriate inflammatory response can lead to a wide range of pathologies including arthritis and cancer. There is evidence to suggest that NO is a key mediator of antiinflammatory processes, by inhibiting the interaction between

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leucocytes and the endothelial monolayer (25). However, both pro- and anti-inflammatory reactions which are independent of soluble guanylate cyclase have been identified (26). The mechanisms by which NO regulates inflammation are not fully understood, but NO is known to inhibit cytokine production, which in turn causes reduced expression of adhesion molecules such as P-selectin (27). eNOS-derived NO also plays an important role in inhibiting thrombogenic events at the blood vessel wall. It has been shown both to inhibit platelet adhesion to the endothelial surface and to reverse aggregation of platelets to vessel wall components (28). It now seems likely that more than one mechanism is involved: cGMP-dependent pathways are certainly implicated, but there is also evidence for cGMP-independent signalling involving S-nitrosylation of proteins (29) and regulation of integrin alpha(IIb)beta (3) and myosin light chain (30). 3.3. NO Generation in Platelets

NO is generated by components of the cardiovascular system other than endothelial cells. It occurs in platelets in response to a wide variety of stimuli including interactions with collagen, von Willebrand factor, β2 agonists, insulin, glucose and vitamin E (31). It has also been suggested that shear stress may induce eNOS in platelets in a manner analogous to that seen in endothelial cells (31). While less efficient in platelets than in endothelial cells, eNOS, nevertheless, activates the gualyate cyclase/cGMP pathway (32, 33). The calcium dependency of eNOS in endothelial cells is well established, but the role of calcium in activation of the NO pathway by collagen in platelets remains somewhat controversial. Activation of cGMP via the collagen glycoprotein receptor has been shown to be calcium dependent (34), but there is also evidence that the NO pathway can be activated in platelets by collagen without an increase in internal calcium concentration (35). This is consistent with the more recent work showing that the role of NO in platelet function is highly complex (34).

3.4. Endothelial Permeability

A key role of the endothelium is to regulate the permeability of the blood vessel wall, particularly in the microvasculature. In common with other aspects of NO biology, concentration is key to determining its effect on NO barrier function. NO at low concentrations can be effective in preventing increases in permeability mediated by components of the coagulation cascade (36), while high concentrations induce nitrosative stress leading to disruption of the endothelial cell barrier (37). In considering the opposing vascular roles of NO at high and low concentrations, it is necessary to include the interaction of NO with reactive oxygen species (ROS) such as hydrogen peroxide (H2 O2 ), super oxide (O2 – ) and hydroxide (OH), because under pathological conditions all may be present to a greater or lesser extent. NO is capable of reacting

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rapidly with damaging radicals, which may result in a reduction in oxidative stress (38) or an increase in nitrosative stress depending on the precise balance of species in the tissue. For example, the rapid reaction of NO with O2 – results in the generation of peroxynitrite (ONOO– ) and this has been shown to degrade the integrity of the endothelial barrier (39). There is also evidence to show that activation of cGMP, a primary target for NO, is cytoprotective for the vessel wall by downregulating cytokines (40), and furthermore NO has been shown to inhibit the activation of mast cells (41). This activity may contribute to the normalising effect of low levels of NO on permeability. At high NO concentrations in the micromolar range, such as those generated in toxic shock by induction of iNOS by bacterial toxins, any cytoprotective effects are overwhelmed by widespread nitrosation of proteins, lipid peroxidation and damage to DNA; this has been shown to result in the activation of apoptosis (42) and depletion of radical scavengers such as GSH (43, 44). 3.5. Angiogenesis

Given its pivotal role in vascular regulation it is not surprising that NO has been identified as a key mediator of angiogenesis. However, its involvement in this process is not simple and it encompasses numerous pathways (45). It was shown over 15 years ago that blockage of NO generation by inhibiting NOS prevented PGE1-induced angiogenesis in the rabbit cornea in vivo and conversely the administration of the NO donor sodium nitroprusside stimulated angiogenesis in these models (46). VEGF-induced differentiation and tubule-forming ability in human endothelial cells (HUVEC) in vitro have also been shown to be NO dependent (47); very similar dependency was also reported for endothelial cells stimulated by bFGF (48) and angiopoietin (49). Thus, the available data indicate that NO is a downstream mediator of the actions of multiple angiogenic effectors. It may, however, also be a reciprocal inducer of VEGF resulting in a mechanism that enhances the strength of angiogenesis. More recently, there has been considerable interest in the asymmetric dimethylarginine (ADMA) pathway and its importance in pathologies resulting from deficiencies in NO generation (50). Furthermore the importance of this molecule in inhibiting various components of angiogenesis has recently been revealed (51). A large body of evidence now suggests that appropriately regulated generation of NO by eNOS is supportive of a healthy cardiovascular function (52). Furthermore, it is widely believed that deficiencies in this NO generating system are a major cause of endothelial dysfunction, resulting in the development of atherosclerosis, hypercholesterolaemia, thrombosis, infarction and stroke (53). Conversely, therapies that deliver NO can protect against cardiovascular disease. Glyceryl trinitrate has been a mainstay of the management of angina for decades (54) and we

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now know that for many of the agents now in clinical use for the treatment of hypercholesterolaemia and cardiovascular disease, enhancement of NO bioavailability is a major component of their actions (55).

4. NO and Injury Repair The healing wound is a highly complex and coordinated system in which NO is a key player (56). It has long been known that L -arginine levels are an important determinant of the effectiveness of healing of skin wounds (57). Initially this was thought to be a consequence of regulation of growth hormone levels by L -arginine, but it was later shown that arginine supplementation was ineffective at enhancing wound healing in animals in which the high output isoform of iNOS had been knocked out, so reinforcing a key role for NO in these processes (58). Arginine enhancement of wound healing has also been demonstrated in healthy human volunteers (59, 60), and NO bioactivity has been proposed as a diagnostic indictor for the management of diabetic ulcers (61). Also, NO-containing nanoparticles have been shown to enhance wound healing (62). The role of NO in the repair of damage in other tissues has also been studied: for example, the process whereby myogenic satellite cells are stimulated to proliferate and to fuse with other myocytes is now known to be regulated by NO together with metalloproteinases and hepatocyte growth factor (63). The correct balance of these factors determines the ability to build functional muscle in response to stress rather than suffer fibrotic scarring, leading to the popularity of arginine supplementation amongst body builders.

5. Nitric Oxide in the Nervous System

The last few years of the 1980s saw a dramatic expansion in our understanding of the roles of NO in mammalian physiology. NO was identified as the “endothelium-derived relaxing factor” (12) and shortly after, it was shown that neurotransmitter responses were mediated by L-arginine-dependent NO generation in neurones (64). It was also shown that some neurons release neither acetylcholine nor catecholamines when stimulated and instead rely upon NO for signal transduction (65). This signalling pathway was also revealed to result in increased cGMP levels (66). NO has been implicated in multiple brain functions including pain perception, memory, thirst and hunger perception and anxiety.

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We now know that NO is a ubiquitous transmitter in both the central and the peripheral nervous systems (67). Peripheral actions have been identified in the penis, where nNOS-containing parasympathetic nerves provide copious innervation to the corpus cavernosum (68, 69) resulting in cGMP-dependent relaxation/expansion (70). The gut is also densely supplied with nitrergic nerves. Numerous studies, mostly using NOS inhibitors, have implicated NO in relaxation of jejunum, colon, rectum and anal sphincter (71), via a cGMP-dependent pathway (72). Most regions of the brain that have been examined express nNOS, but the vasculature that supplies the brain is also innervated by neurons expressing NO as well as acetylcholine (67) and these regulate vascular tone via c-GMP dependent mechanisms (73). A key function of NO in this system is to modulate cerebral blood flow in response to changes in blood gasses O2 and CO2 , and as such it plays a vital role in regulating substrates for brain metabolism (74). It is not surprising, therefore, that deficiencies in the regulation of NO generation by these nerves can lead to a number of cerebral pathologies including migraine, inflammatory disorders and even Alzheimer’s and Huntington’s disease. It may also mediate the damaging effects of ethanol in the central nervous system (75). Within the reticular activating system NO appears to play an important role in modulating overall brain activity; in particular, NO levels in the thalamus may determine wakefulness and sleep patterns (76, 77).

6. NO and Addiction There is now clear evidence to support the view that the rewarding effects of addictive substances in the brain are mediated at least in part by NO (78). Most of this information has been obtained in animal studies using reward/preference testing with and without the administration of NOS inhibitors. In the case of ethanol, NOS inhibition reduced preference/consumption and reduced withdrawal in rats (79, 80), but enhanced the acute central depressant and anaesthetic effects of alcohol (81). NO is also a key modulator of responses to nicotine and both peripheral and central effects are involved (82), though the responses to smoking activities are complicated by the fact that NO is a significant constituent of tobacco smoke –approximately 200 µg/cigarette (83). It also causes bronchodilation, allowing smoke to penetrate the lungs more effectively and reduces pulmonary vascular resistance (84, 85). Key to its role in the psychoactive effects of nicotine is the observation that NO enhances the dopamine-dependent reward system (86, 87) that is thought to be fundamental to the addictive nature of most drugs of dependency including nicotine

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(88). There is also evidence that the behavioural effects resulting from addiction to opiates and cocaine (89–91) as well as to ecstasy and chloroamphetamine (92) are mediated by NO and that NOS inhibitors reduce sensitivity to opiates (93). Given the ubiquitous involvement of NO in the reward mechanism through dopamine enhancement, it is reasonable to speculate that NO may contribute to some positive effects of placebos (94).

7. NO and Cancer NO is also generated at higher concentrations in tumours than in most normal tissues of origin (6). There is now compelling evidence that NO generated by all NOS isoforms plays major roles in the development of cancer and in the response of tumours to most therapies. It is also believed that eNOS may be key to the numerous processes that are characteristic of the malignant phenotype (95). Oncogenesis in many tissues is driven by chronic inflammation (96) and it is recognized that NO can regulate proinflammatory mediators leading to oncogenic transformation (97). Another key determinant of malignant progression is apoptosis and the concentration of NO has been shown to influence the balance between proliferation and cell death (98). The action of NO on the tumour vasculature also determines clinically relevant processes in tumours; excessive NO production in solid tumours has been shown to increase vascular permeability and blood flow, thus promoting tumour growth and the efficiency of metastatic spread (99). It is also a key regulator of angiogenesis (100), which is fundamental to the ability of cancer cells to attract a blood supply and then to disseminate to distant sites. Even a concise summary of the roles of NO in cancer reveals a complex picture (95). Nevertheless, NO has been identified as an important target for cancer therapy and as a therapeutic agent in its own right (101). Therapies involving the delivery of high concentrations of NO via donor drugs or gene therapy have proven to be effective against experimental cancer models (6, 102) and early clinical trials in lung (103) and prostate cancer (104) have been impressive.

8. Nitric Oxide in Non-mammalian Systems

Far from being the preserve of higher vertebrates, NO generated from NOSs is a highly conserved system that has been found in most species that have been examined. Indeed it is difficult to find

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organisms that do not express a form of NOS, Caenorhabditis elegans being a distinct anomaly (105) that expresses guanylate cyclase isoforms that are insensitive to NO (106). NOS isoforms in insects, molluscs, crustaceans and other invertebrates have been well characterized (67). Nitric oxide is now known to be as important in plants as it is in animals (107). It was first shown to have a major role in disease defence (108), but it has since been shown to be vital for fundamental processes such as seed germination and flower and root development (109). NOSs are even expressed in bacteria where they often lack reductase domains and require additional reductants to generate NO. Bacterial NOSs have functions that differ from those isoforms found in vertebrates, including protection against oxidative stress (110). Thus, while NO has been a ubiquitous regulator of cell signalling in living organisms for millions of years, the detailed understanding of its role in physiology now provides us with the opportunity to harness it for human benefit. References 1. Wang, R. (2002) Two’s company, three’s a crowd: can H2S be the third endogenous gaseous transmitter? FASEB J 16, 1792–1798. 2. Mustafa, A. K., Gadalla, M. M., Snyder, S. H. (2009) Signaling by gasotransmitters. Sci Signal 2, 1–8. 3. Aranda, M., Pearl, R. G. (2000) Inhaled nitric oxide and pulmonary vasoreactivity. J Clin Monit Comput 16, 393–401. 4. Iyengar, R., Stuehr, D. J., Marletta, M. A. (1987) Macrophage synthesis of nitrite, nitrate, and N-nitrosamines: precursors and role of the respiratory burst. Proc Natl Acad Sci USA 84, 6369–6373. 5. Hibbs, J. B., Jr., Taintor, R. R., Vavrin, Z., Rachlin, E. M. (1988) Nitric oxide: a cytotoxic activate macrophage effector molecule. Biochem Biophys Res Commun 157, 87–94. 6. Hirst, D., Robson, T. (2007) Targeting nitric oxide for cancer therapy. J Pharm Pharmacol 59, 3–13. 7. Xu, W., Charles, I. G., Moncada, S. (2005) Nitric oxide: orchestrating hypoxia regulation through mitochondrial respiration and the endoplasmic reticulum stress response. Cell Res 15, 63–65. 8. Brown, G. C. (2001) Regulation of mitochondrial respiration by nitric oxide inhibition of cytochrome c oxidase. Biochim Biophys Acta 1504, 46–57. 9. Aguirre, E., Rodriguez-Juarez, F., Bellelli, A., Gnaiger, E., Cadenas, S. (2010) Kinetic

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65. Bult, H., Boeckxstaens, G. E., Pelckmans, P. A., Jordaens, F. H., Van Maercke, Y. M., Herman, A. G. (1990) Nitric oxide as an inhibitory non-adrenergic non-cholinergic neurotransmitter. Nature 345, 346–347. 66. Bredt, D. S., Snyder, S. H. (1989) Nitric oxide mediates glutamate-linked enhancement of cGMP levels in the cerebellum. Proc Natl Acad Sci USA 86, 9030–9033. 67. Vincent, S. R. (2010) Nitric oxide neurons and neurotransmission. Prog Neurobiol 90, 246–255. 68. Rajfer, J., Aronson, W. J., Bush, P. A., Dorey, F. J., Ignarro, L. J. (1992) Nitric oxide as a mediator of relaxation of the corpus cavernosum in response to nonadrenergic, noncholinergic neurotransmission. N Engl J Med 326, 90–94. 69. Burnett, A. L., Lowenstein, C. J., Bredt, D. S., Chang, T. S., Snyder, S. H. (1992) Nitric oxide: a physiologic mediator of penile erection. Science 257, 401–403. 70. Recio, P., Lopez, P. G., Hernandez, M., Prieto, D., Contreras, J., Garcia-Sacristan, A. (1998) Nitrergic relaxation of the horse corpus cavernosum. Role of cGMP. Eur J Pharmacol 351, 85–94. 71. Rolle, U., Nemeth, L., Puri, P. (2002) Nitrergic innervation of the normal gut and in motility disorders of childhood. J Pediatr Surg 37, 551–567. 72. Shuttleworth, C. W., Xue, C., Ward, S. M., de Vente, J., Sanders, K. M. (1993) Immunohistochemical localization of 3, 5-cyclic guanosine monophosphate in the canine proximal colon: responses to nitric oxide and electrical stimulation of enteric inhibitory neurons. Neuroscience 56, 513–522. 73. Gonzalez, C., Barroso, C., Martin, C., Gulbenkian, S., Estrada, C. (1997) Neuronal nitric oxide synthase activation by vasoactive intestinal peptide in bovine cerebral arteries. J Cereb Blood Flow Metab 17, 977–984. 74. Toda, N., Ayajikim, K., Okamura, T. (2009) Cerebral blood flow regulation by nitric oxide: the recent advances. Pharmacol Rev 61, 62–97. 75. Lancaster, F. E. (1995) Alcohol and the brain: what’s NO got to do with it? Metab Brain Dis 10, 125–133. 76. Williams, J. A., Vincent, S. R., Reiner, P. B. (1997) Nitric oxide production in rat thalamus changes with behavioral state, local depolarization, and brainstem stimulation. J Neurosci 17, 420–427. 77. Hars, B. (1999) Endogenous nitric oxide in the rat pons promotes sleep. Brain Res 816, 209–219.

78. Tayfun Uzbay, I., Oglesby, M. W. (2001) Nitric oxide and substance dependence. Neurosci Biobehav Rev 25, 43–52. 79. Rezvani, A. H., Grady, D. R., Peek, A. E., Pucilowski, O. (1995) Inhibition of nitric oxide synthesis attenuates alcohol consumption in two strains of alcoholpreferring rats. Pharmacol Biochem Behav 50, 265–270. 80. Lallemand, F., De Witte, P. (1997) LNNA decreases cortical vascularization, alcohol preference and withdrawal in alcoholic rats. Pharmacol Biochem Behav 58, 753–761. 81. Adams, M. L., Cicero, T. J. (1998) Alcohol intoxication and withdrawal: the role of nitric oxide. Alcohol 16, 153–158. 82. Vleeming, W., Rambali, B., Opperhuizen, A. (2002) The role of nitric oxide in cigarette smoking and nicotine addiction. Nicotine Tob Res 4, 341–348. 83. Liu, C., Feng, S., van Heemst, J., McAdam, K. G. (2010) New insights into the formation of volatile compounds in mainstream cigarette smoke. Anal Bioanal Chem 396, 5, 1817–1830. 84. Vleeming, W., Rambali, B., Opperhuizen, A. (2002 Aug) The role of nitric oxide in cigarette smoking and nicotine addiction. Nicotine Tob Res 4, 3, 341–348. 85. Thebaud, B., Arnal, J. F., Mercier, J. C., Dinh-Xuan, A. T. (1999 Jul) Inhaled and exhaled nitric oxide. Cell Mol Life Sci 55, 1103–1112. 86. Pogun, S., Kuhar, M. J. (1994) Regulation of neurotransmitter reuptake by nitric oxide. Ann N Y Acad Sci 738, 305–315. 87. Dhir, A., Kulkarni, S. K. (2007) Involvement of nitric oxide (NO) signaling pathway in the antidepressant action of bupropion, a dopamine reuptake inhibitor. Eur J Pharmacol 568, 177–185. 88. Govind, A. P., Vezina, P., Green, W. N. (2009) Nicotine-induced upregulation of nicotinic receptors: underlying mechanisms and relevance to nicotine addiction. Biochem Pharmacol 78, 756–765. 89. Itzhak, Y., Martin, J. L., Black, M. D., Huang, P. L. (1998) The role of neuronal nitric oxide synthase in cocaine-induced conditioned place preference. Neuroreport 9, 2485–2488. 90. Manzaned, C., Aguilar, M. A., Do Couto, B. R., Rodriguez-Arias, M., Minarro, J. (2009) Involvement of nitric oxide synthesis in sensitization to the rewarding effects of morphine. Neurosci Lett 464, 67–70. 91. Zarrindast, M. R., Karami, M., Sepehri, H., Sahraei, H. (2002) Influence of nitric

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Chapter 2 Ionizing Radiation-Induced DNA Strand Breaks and γ-H2AX Foci in Cells Exposed to Nitric Oxide Kai Rothkamm and Susanne Burdak-Rothkamm Abstract A number of studies have demonstrated that nitric oxide enhances radiosensitivity of anoxic and hypoxic cells in vitro and in vivo, and some evidence points to a role for DNA damage and repair in this phenomenon. We have recently observed that nitric oxide enhances the formation of DNA single- and double-strand breaks following ionising irradiation, measured by the alkaline comet assay and immunofluorescence microscopy for γ-H2AX. Key words: Nitric oxide, DNA strand break, γ-H2AX, single cell gel electrophoresis, ionising radiation.

1. Introduction Whilst numerous studies have clearly demonstrated that nitric oxide (NO) enhances radiosensitivity of anoxic and hypoxic cells in vitro and in vivo (1, 2), the exact mechanisms underlying the observed radiosensitisation remain elusive. NO has been reported to upregulate p53, PARP and the catalytic subunit of DNA-dependent protein kinase (DNA-PKcs), an enzyme involved in repairing DNA double-strand breaks via the non-homologous end-joining pathway (3–5). On the other hand, it has been reported to inhibit nucleotide excision repair (6). However, it is possible that these data were confounded by the presence of nitrogen dioxide (NO2 ). Furthermore, there is some evidence for the involvement of NO in ‘bystander’ responses (7) which may occur via the radiation-induced activation of nitric oxide synthase (NOS) (8). H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_2, © Springer Science+Business Media, LLC 2011

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Some recent evidence suggests a modulation of radiationinduced DNA damage and/or its repair by NO which appears to correlate with potent low-dose radiosensitisation of cell cultures at low concentrations of NO (9). Ionising radiation induces a wide spectrum of different DNA lesions, ranging from damaged bases and adducts to cross-links and strand breaks. However, few assays are available to measure the formation and repair of DNA damage following low-dose irradiation. We have observed the yield of DNA double-strand breaks in V79 Chinese hamster lung fibroblasts and MCF-7 human breast cancer cells, as detected by immunofluorescence microscopy for the phosphorylated histone variant γ-H2AX. Double-strand breaks increased twofold following X-ray-irradiation in the presence vs. absence of 1% v/v NO in nitrogen (N2 ), and repair time was longer in cells irradiated in NO than in air or N2 alone. Also, single-strand breaks, detected by alkaline single cell gel electrophoresis (‘comet’ assay), appeared to be enhanced in the presence of NO. Furthermore, loss of Xray-induced γ-H2AX foci appeared to be slower in cells exposed to NO (9). These methods are described below.

2. Materials 2.1. Cell Culture, NO Exposure and X-Irradiation

1. Eagle’s MEM with Earle’s salts supplemented with 10% (V79-379A) or 15% (MCF-7) foetal calf serum, penicillin, streptomycin, glutamine, sodium bicarbonate, sodium pyruvate and non-essential amino acids. 2. Solution of 0.25% trypsin and 1 mM ethylendiamine tetraacetic acid (EDTA). 3. NO (100% or 1% v/v in N2 ; 400 ppm v/v in N2 ‘INOmax’ from INO Therapeutics, Sittingbourne, Kent, UK) and N2 . 4. Tubing, valves and fittings made of stainless steel, glass or PEEK polymer; glass syringes for irradiation of pre-gassed cell samples; glass scintillation vials with SubaSeal stoppers and gas entry and exit needles for irradiation of continuously gassed cell samples (see Note 1). 5. X-ray generator (Pantak, East Haven, CT, USA) with 4.3 mm aluminium filtration at 240 kV, 13 mA and a dose rate of approximately 0.5 Gy/min.

2.2. Immunofluorescence for γ-H2AX

1. Lab-Tek II chamber slides and coverslips (22 mm × 50 mm × 0.13 mm). 2. Phosphate-buffered saline (prepared from PBS tablets). 3. 100% methanol (stored at –20◦ C).

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4. Mouse monoclonal antibody (clone JBW301) for γ-H2AX from Millipore, Watford, Hertfordshire, UK (see Note 2). Stable at –20◦ C for several years. 5. AlexaFluor 488-conjugated goat anti-mouse IgG secondary antibody. Stable at 4◦ C for 1 year. Protect from light. 6. 4,6-diamidino-2-phenylindole (DAPI) at a final concentration of 0.5 µg/mL. Stable at –20◦ C for several years. 7. Mounting medium containing TASHIELD. Store at 4◦ C.

antifade,

e.g.

VEC-

8. Clear, non-fluorescent nail varnish. 2.3. Alkaline Single Cell Gel Electrophoresis for DNA Strand Breaks

1. Glass slides and coverslips. 2. Lysis buffer (1 l): 1.2 M NaCl, 0.1% N-lauryl sarcosine, 0.26 M NaOH, 100 mM Na2 EDTA. pH>12.5. 3. Electrophoresis buffer (2 L): 0.03 M NaOH, 2 mM Na2 EDTA, pH>12.5. (The buffers can be made the day before and kept at 4◦ C but check pH before use.) 4. Rinse buffer: previously used electrophoresis buffer. 5. 1% pulsed-field certified agarose and 1% Type VII (low melting point) agarose. 6. Metal plate. 7. Horizontal electrophoresis chamber and power supply. 8. 70, 90 and 100% ethanol. 9. Sybr-Gold DNA staining solution. Store at –20◦ C, protected from light. 10. DABCO antifade mounting medium. Store at –20◦ C. 11. Clear nail varnish.

3. Methods Following the induction of a DNA double-strand break, hundreds to thousands of copies of the histone variant H2AX, which forms part of the nucleosome, are phosphorylated by the DNA damageactivated kinases, Ataxia Telangiectasia Mutated (ATM) and DNA-dependent Protein Kinase (DNA-PK), covering megabases of chromatin surrounding the site of the break. Immunofluorescence microscopy using commercially available antibodies for the phosphorylated form, γ-H2AX, can be employed to visualise and quantify these radiation-induced ‘foci’ which are used

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as a surrogate marker for double-strand breaks and their repair (10, 11). The comet assay detects DNA damage in single cells following gel electrophoresis of low concentrations of lysed cells embedded in agarose (12). The type of DNA damage detected in the comet assay depends primarily on the pH of buffers used for cell lysis and electrophoresis. Neutral conditions are used to measure double-stranded DNA breaks, whereas alkaline conditions allow the detection of both single- and double-stranded DNA breaks, as the alkaline conditions lead to denaturation of DNA. Given that ionising radiation induces 20–50 times more single- than doublestrand breaks, most of the damage measured in the alkaline comet assay reflects single-strand breaks. DNA damage is induced immediately during irradiation, and for determining initial yields of DNA damage, cells should be put on ice immediately after irradiation to suppress repair. This is especially important for DNA strand breaks measured with the alkaline comet assay because of the rapid repair of DNA single-strand breaks. However, for γ-H2AX-based measurements of DNA double-strand breaks, cells require post-irradiation incubation at 37◦ C to enable efficient phosphorylation of H2AX at sites of double-strand breaks. Depending on the quality of the immunofluorescence staining and on the background levels present in different cell lines, minimum incubation times of 3–30 min are commonly used, and, accordingly, γ-H2AX levels never reflect the full initial yield of damage (13). In the experiments described below, cells are irradiated in suspension and transferred to chamber slides. In this case the minimum incubation time is 30 min at 37◦ C to allow sufficient numbers of cells to attach to the slide surface. 3.1. Cell Culture, Nitric Oxide Exposure and X-RayIrradiation

1. V79 and MCF-7 cells are split with trypsin/EDTA when approaching confluence and passaged in T25 tissue culture flasks to maintain cultures. Doubling times are 10–12 h for V79 and approximately 20 h for MCF-7 cells. 2. To prevent NO autoxidation, suspensions of exponentially growing cells (∼7 × 105 in PBS) are first pre-gassed by gently bubbling with argon (Ar) or N2 for 30 min before gassing for 30 min with the appropriate gas (see Note 3). 3. All irradiations are performed at room temperature. Either the suspensions are irradiated in capped glass syringes and 1 mL samples are collected after each X-ray dose and placed on ice or 2 mL suspensions are irradiated in glass scintillation vials held nearly horizontal (5–10◦ angle with ∼22 mL headspace) whilst being continuously bubbled with the appropriate gas. The vials are placed on ice after exposure (see Note 4).

DNA Strand Breaks and γ-H2AX Foci in Cells Exposed to NO

3.2. Immunofluorescence for γ-H2AX

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1. The cell suspensions are transferred into Lab-Tek II chamber slides and incubated at 37◦ C in a humid atmosphere containing 5% CO2 . 2. Cells are fixed with 100% methanol at –20◦ C for 10 min (see Note 5). 3. Cells are incubated in PBS with 2% foetal calf serum for 3 × 5–10 min incubations at room temperature to block non-specific antibody binding sites (see Note 6). 4. Samples are incubated with anti γ-H2AX antibody (1:300 in PBS, 2% foetal calf serum) for 1 h at room temperature. 5. Samples are washed in PBS with 2% foetal calf serum for 3 × 5–10 min washes at room temperature. 6. Samples are incubated with secondary antibody (1:400 in PBS, 2% foetal calf serum) for 1 h at room temperature in the dark. 7. Slides are washed in PBS for 5–10 min at room temperature in the dark. 8. Cells are counterstained with DAPI for 3–5 min and washed in PBS for 5–10 min at room temperature in the dark. 9. Chambers are removed from slides and slides completely dried in the dark before applying mounting medium and mounting with a coverslip. Nail varnish is applied to seal the samples. Samples can be viewed when the varnish is dry and can be stored in the dark at 4◦ C for several weeks. 10. The slides are viewed using a fluorescence microscope. An example is shown in Fig. 2.1. Scoring of nuclear γ-H2AX foci typically requires at least an ×40 objective and can be

Fig. 2.1. Foci of γ-H2AX after irradiation of anoxic V79 cells in sealed syringes following incubation for 1 h at 37◦ C in 90/5/5% v/v N2 /O2 /CO2 . Upper panel: irradiated with 0, 0.2, 0.4 and 1 Gy in N2 ; lower panel: irradiated with 0, 0.2, 0.4 and 1 Gy in 1% v/v NO in N2 .

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performed by eye either through the microscope eye pieces or on images obtained with a camera. Alternatively, a range of software packages have been used to facilitate automated scoring of γ-H2AX foci (14–17) (see Note 7).

4. Alkaline Single Cell Gel Electrophoresis for DNA Strand Breaks

1. Furnace BDH semi-frosted slides. 2. Coat non-frosted part of slides with 100 µL of 1% PFGE agarose, air-dry slides and store in 50◦ C oven. 3. Label slides coated earlier using a pencil. 4. Make up 1% Type VII agarose and place in water bath at 37◦ C until needed. 5. Place metal plate in freezer. 6. Count cells and dilute to 50,000/mL, using cold medium. 7. Remove metal plate from freezer and cover with four sheets of paper towel. 8. For each slide aliquot 125 µL cell suspension in 5 mL tube and keep on ice. Add 375 µL of 1% Type VII agarose to cell suspension, gently mix and spread quickly on a coated slide. Place on metal plate and allow to set. Place in lysis buffer in a container in fridge and lyse overnight. 9. Carefully remove slides from lysis buffer and place slides in 650 mL of rinse buffer for 20 min, repeat twice. 10. Fill the electrophoresis tank with 1.6 L electrophoresis buffer. Put slides on plastic tray in electrophoresis tank (ensure that the slides are straight and all facing in the same direction). Run at 0.6 V/cm for 30 min. When the run ends, immediately remove the slides from the tank and place in cold 70% EtOH for 10 min, followed by 90% EtOH for 10 min and 100% EtOH for 10 min. Air-dry. The slides are now stable and can be left for any period of time before proceeding with the staining. 11. Make up a 1:10,000 dilution of Sybr-Gold in 1 × TBS. Sybr-Gold is light sensitive and must be kept in the dark. Incubate with slides for 30 min in the dark. Rinse twice for 5 min with distilled H2 O, remove the slides and drain, but do not allow to dry out. 12. Put 50 µL DABCO antifade along the centre of a coverslip. Lower the slide onto it and gently push down, leave for a few minutes to settle, then remove any excess. Seal around the edges of the coverslip with nail varnish. Leave to dry.

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13. Slides are analysed either by an automated microscope image acquisition and analysis system which allows scoring of thousands of cells per hour or by manual analysis using a fluorescence microscope (see Note 8).

5. Notes 1. Standard tissue culture plasticware and plastic tubing should be avoided as it contains oxygen which can leak out from the surface and thus contaminate the sample. 2. We have found this antibody to work very reliably for immunofluorescence. Numerous polyclonal rabbit antibodies for γ-H2AX are available from other commercial suppliers but have suffered from excessive non-specific staining and high variability in our hands. 3. Removal of NO2 can be confirmed by the failure of the purified NO to oxidise N2-purged aqueous solutions of the dye ABTS [2,2′ -azino-bis(3-ethylbenzothiazoline-6-sulfonate)] to its stable radical (green colour). 4. Continuous bubbling during irradiation ensures that NO levels are not diminished during the irradiation process. This is especially important at low concentrations of NO and at high radiation doses. 5. Fixation may have to be adjusted, depending on the cell line used. If the DAPI signal shows that DNA seems to be leaking out of the nuclei, then this indicates that the coarse precipitation of proteins facilitated by methanol treatment has broken up cell nuclei quite severely. Such disruption of cellular morphology occurs frequently in some cell types, e.g. leukocytes, when using methanol for fixation. It tends also to ‘smear out’ gamma-H2AX foci which, as a result, are more difficult to discern and score. To improve the preservation of cell morphology, cells can be pre-fixed with 2–4% buffered formaldehyde (prepared fresh in PBS from a buffered stock of 37% formaldehyde solution or from paraformaldehyde) for about 10 min at room temperature before a 10 min methanol fixation step at –20◦ C. The protein and DNA cross-links resulting from this treatment stabilise cellular structures and thereby help obtain more distinct gamma-H2AX foci. 6. The blocking efficiency of different batches of foetal calf serum may vary. Bovine serum albumin (fraction V) or other sera may be used as an alternative. This step and/or the subsequent incubation with primary antibody can be extended to overnight incubation at 4◦ C, if required or convenient.

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7. In order to obtain the total number of foci per cell, nuclei must be imaged across their depth, and not just a single image taken. For software-based scoring, either 3D image analysis software or 2D analysis of maximum projections of z-stacks of images should be used. The required step size for image stacks depends on the focus depth of the optical system and is typically in the range of 200–1,000 nm. With increasing foci levels, more and more overlap between adjacent foci occurs which may result in ‘underscoring’. The extent of this effect depends on the optical resolution of the system and, for automated scoring, on the software algorithms used to detect and distinguish individual foci. Optionally, 3D deconvolution or confocal microscopy can be used to enhance the optical resolution along the depth axis. In practice, however, these rather time-consuming and expensive efforts rarely improve the accuracy of foci quantification. 8. A comet consists of a ‘head’ of DNA which cannot migrate out of the nucleus due to its size, and a ‘tail’ of fragmented DNA leaving the nucleus during electrophoresis and migrating according to the molecule size. The ‘tail length’ (distance between the centres of mass of the tail and head region), % of DNA in the tail and ‘tail moment’ (the product of tail length and % DNA in the tail) can be measured and calculated using dedicated image analysis software, and these terms are used to describe and quantify the amount of DNA damage measured. Alternatively, if analysing comets by eye, they can be classified into different categories of damage, depending on their appearance. See (18) for a more detailed discussion of analysis techniques and statistical issues in the use of the comet assay.

Acknowledgements The methods described here were employed in a recent Cancer Research UK-funded study at the Gray Cancer Institute under Peter Wardman’s leadership, with support from Mick Woodcock, Lisa Folkes and Peter Johnston. References 1. Wardman, P. (2007) Chemical radiosensitizers for use in radiotherapy. Clin Oncol (R Coll Radiol) 19, 397–417. 2. Hirst, D. G., Robson, T. (2007) Nitrosative stress in cancer therapy. Front Biosci 12, 3406–3418.

3. Xu, W., Liu, L. Z., Loizidou, M., Ahmed, M., Charles, I. G. (2002) The role of nitric oxide in cancer. Cell Res 12, 311–320. 4. Xu, W., Liu, L., Smith, G. C., Charles, L. G. (2000) Nitric oxide upregulates expression of DNA-PKcs to protect cells from

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6.

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8.

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11.

DNA-damaging anti-tumour agents. Nat Cell Biol 2, 339–345. Cook, T., Wang, Z., Alber, S., Liu, K., Watkins, S. C., Vodovotz, Y., Billiar, T. R., Blumberg, D. (2004) Nitric oxide and ionizing radiation synergistically promote apoptosis and growth inhibition of cancer by activating p53. Cancer Res 64, 8015–8021. Chien, Y., Bau, D., Jan, K. (2004) Nitric oxide inhibits DNA-adduct excision in nucleotide excision repair. Free Radic Biol Med 36, 1011–1017. Shao, C., Folkard, M., Michael, B. D., Prise, K. M. (2004) Targeted cytoplasmic irradiation induces bystander responses. Proc Natl Acad Sci USA 101, 13495–13500. Leach, J. K., Black, S. M., Schmidt-Ullrich, R. K., Mikkelsen, R. B. (2002) Activation of constitutive nitric-oxide synthase activity is an early signaling event induced by ionizing radiation. J Biol Chem 277, 15400–15406. Wardman, P., Rothkamm, K., Folkes, L. K., Woodcock, M., Johnston, P. J. (2007) Radiosensitization by nitric oxide at low radiation doses. Radiat Res 167, 475–484. Rothkamm, K., Krüger, I., Thompson, L. H., Löbrich, M. (2003) Pathways of DNA double-strand break repair during the mammalian cell cycle. Mol Cell Biol 23, 5706–5715. Rogakou, E., Boon, C., Redon, C., Bonner, W. (1999) Megabase chromatin domains

12.

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involved in DNA double-strand breaks in vivo. J Cell Biol 146, 905–916. McKenna, D. J., McKeown, S. R., McKelveyMartin, V. J. (2008) Potential use of the comet assay in the clinical management of cancer. Mutagenesis 23, 183–190. Rothkamm, K., Horn, S. (2009) GammaH2AX as protein biomarker for radiation exposure. Ann Ist Super Sanita 45, 265–271. Qvarnstrom, O. F., Simonsson, M., Johansson, K. A., Nyman, J., Turesson, I. (2004) DNA double strand break quantification in skin biopsies. Radiother Oncol 72, 311–317. Barber, P. R., Locke, R. J., Pierce, G. P., Rothkamm, K., Vojnovic, B. (2007) GammaH2AX foci counting: image processing and control software for high content screening. Proc SPIE 6441, M1–M10. Costes, S. V., Boissiere, A., Ravani, S., Romano, R., Parvin, B., Barcellos-Hoff, M. H. (2006) Imaging features that discriminate between foci induced by high- and lowLET radiation in human fibroblasts. Radiat Res 165, 505–515. Bocker, W., Iliakis, G. (2006) Computational methods for analysis of foci: validation for radiation-induced gamma-H2AX foci in human cells. Radiat Res 165, 113–124. Lovell, D. P., Omori, T. (2008) Statistical issues in the use of the comet assay. Mutagenesis 23, 171–182.

Chapter 3 Determination of S-Nitrosothiols in Biological Fluids by Chemiluminescence Enika Nagababu and Joseph M. Rifkind Abstract S-nitrosothiols present in nanomolar concentrations in cells and body fluids play an important role in vasodilation, in preventing platelet aggregation, leukocyte adhesion, and for cellular signaling. However, because of the low levels of s-nitrosothiols and interference with other nitric oxide species, reliable assays that measure both high molecular weight and low molecular weight s-nitrosothiols in plasma and red blood cells have been difficult to develop. We have previously developed a sensitive method using Cu(II)-ascorbic acid at a neutral pH, which was specific for s-nitrosothiols without interference of nitrite or other NOx species. However, due to neutral pH foaming, this method was not suitable for determinations in plasma or red blood cells with high protein content. This method has now been modified by using copper (II) chloride (CuCl2 ) and ascorbic acid in glacial acetic acid. The low pH solves the foaming problem. However, protonation of nitrite under acidic conditions facilitates the formation of s-nitrosothiols. For this method to specifically measure s-nitrosothiols in the sample, the unreacted thiols are blocked by reacting with N-ethylmaleimide and nitrite is blocked by reacting with acidified sulfanilamide before being analyzed by chemiluminescence. Using this method, s-nitrosothiols have been determined in the range of 2 nM to 26 nM (mean ± SE = 10.18±2.1) in plasma and up to 88.1 nM (mean ± SE = 51.27 ± 10.5) in red blood cells. Key words: S-nitrosothiols, nitric oxide, ozone-based chemiluminescence assay, plasma, red blood cells.

1. Introduction S-nitrosothiols (RSNOs) are adducts of nitric oxide (NO) with thiol groups. NO does not directly react with thiols to form RSNOs at physiological pH (1). However, oxidation of NO by Research Support: This research was supported by the Intramural Research Program of the NIH, National Institute on Aging

H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_3, © Springer Science+Business Media, LLC 2011

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oxygen forms dinitrogen trioxide, which can react with thiols to form RSNOs (2). High molecular weight and low molecular weight RSNOs are present in nanomolar concentrations in vivo in cells and body fluids. It has been proposed that RSNOs in the plasma play a role in vasodilation, antiplatelet aggregation activity, and anti-leukocyte adhesion properties (3, 4). They also serve as intracellular signaling molecules (5). Blood RSNO levels have been shown to vary under various pathological conditions and therefore have potential clinical relevance (6). Determination of total or individual S-nitrosylated proteins is very challenging. Several methods have been developed to determine the total RSNOs in plasma, red blood cells, and whole blood (7). The most popular and simple method for the determination of RSNOs is the Saville reaction involving the treatment of RSNOs with mercuric chloride, which releases NO+ that then reacts with Griess reagents to form azodye which can be detected colorimetrically. Fluorescence methods have also been developed for the detection of RSNOs using 4, 5-diaminofluorescein dyes. However, these methods are not sensitive enough to measure the physiological levels of RSNOs that are expected to be 18 M cm), to avoid contamination and consequent interference by trace metals. 7. This solution should be stored at 4◦ C and protected from light by a foil wrapper. It should be discarded after 1 week. 8. This solution should be prepared every day and protected from light by a foil wrapper. It can be used for at least 8 h. 9. Piston pumps equipped with syringes or current liquid chromatography pumps are also applicable. 10. Software is commercially available or it can be created from R programming tools such as Visual Basic or Lab View . 11. After assembling the system, check that there are no leaks by performing the procedure using H2 O instead of reagents. 12. Make sure that the end of the synthesis coil is dipped in the NaOH solution. 13. The ONOO– stock solution may be frozen at –80◦ C. ONOO– gradually decomposes with a half-life of 1–2 weeks, yielding nitrite. Storage at –20◦ C can result in an increased concentration of ONOO– as a supernatant layer is formed above the ice crystals. At this layer ONOO– concentration can reach up to 1 M. 14. Dilute the ONOO– stock solution 100–1,000-fold, using 1.2 M NaOH as the diluent in order to obtain an absorbance value within the linear Lambert–Beer relation. 15. The connection between the detector and the confluence where luminol is added must be as short as possible because light production is fast. 16. Check that solutions are aspirated by removing the pumping tubing a few times from the solution, allowing air bubbles to enter the tube. Observe the movement of air bubbles along the system and if flow pulses (short stops of flow) exist, change the pressure exerted from the pump braces so

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that no pulses are visible. If any of the solutions are not aspirated, check if the pump braces are correctly adjusted. 17. Discard or filter solutions containing particulates as they can block the flow tubing. 18. Check the flow rate by placing the pump tubing in a measuring cylinder filled with H2 O. Determine the volume of H2 O aspirated during 1 min, which will correspond to the flow rate expressed as millilitre per minute. 19. If necessary, adjust the flow rate by changing the rotation speed of the pump. 20. To fill the injection loop, the test solution can be aspirated through extra pump tubing placed in the peristaltic pump or it can be aspirated by an external syringe. 21. For positive controls, CL quenching should be observed in a concentration-dependent manner. For negative controls, no CL quenching is observed. 22. It is necessary to stop the pump while the port position is changed at the selection valve. This can be easily achieved by computer-controlled operation of equipment. 23. It is essential to perform blank measurements (using sample diluent as test solution) at the beginning and end of analysis. Blank signal peak height is necessary to calculate the inhibition percentage. Solution stability can be assessed by the reproducibility of blank measurement peak height over time. 24. If organic solvent interference is verified, replace the H2 O by a solution with the same composition of the samples (e.g., same organic solvent percentage) as carrier solution. Analyze positive and negative controls to evaluate the performance of the method. References 1. Ruzicka, J., Hansen, E. H. (1975) Flow injection analyses. 1. New concept of fast continuous-flow analysis. Anal Chim Acta 78, 145–157. 2. Hansen, E. H., Norgaard, L., Pedersen, M. (1991) Optimization of flow-injection systems for determination of substrates by means of enzyme amplification reactions and chemiluminescence detection. Talanta 38, 275–282. 3. Ruzicka, J., Marshall, G. D. (1990) Sequential injection – a new concept for chemical sensors, process analysis and laboratory assays. Anal Chim Acta 237, 329–343.

4. Segundo, M. A., Magalhães, L. M. (2006) Multisyringe flow injection analysis: stateof-the-art and perspectives. Anal Sci 22, 3–8. 5. Magalhães, L. M., Santos, M., Segundo, M. A., Reis, S., Lima, J. L. F. C. (2009) Flow injection based methods for fast screening of antioxidant capacity. Talanta 77, 1559–1566. 6. Magalhães, L. M., Lucio, M., Segundo, M. A., Reis, S., Lima, J. L. F. C. (2009) Automatic flow injection based methodologies for determination of scavenging capacity against biologically relevant reactive species of oxygen and nitrogen. Talanta 78, 1219–1226.

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7. Magalhães, L. M., Ribeiro, J. P. N., Segundo, M. A., Reis, S., Lima, J. L. F. C. (2009) Multi-syringe flow-injection systems improve antioxidant assessment. Trend Anal Chem 28, 952–960. 8. Sariahmetoglu, M., Wheatley, R. A., Cakici, I., Kanzik, I., Townshend, A. (2003) Flow injection analysis for monitoring antioxidant effects on luminol chemiluminescence of reactive oxygen species. Anal Lett 36, 749–765. 9. Beckman, J. S., Chen, J., Ischiropoulos, H., Crow, J. P. (1996) Oxidative chemistry of peroxynitrite. Method Enzymol 269, 229–240. 10. Yao, D. C., Vlessidis, A. G., Evmiridis, N. P. (2001) On-line monitoring of nitric oxide complexed with porphyrine-bearing biochemical materials by using flow injection with chemiluminescence detection. Anal Chim Acta 435, 273–280. 11. Yao, D. C., Vlessidis, A. G., Evmiridis, N. P., Evangelou, A., Karkabounas, S., Tsampalas, S. (2002) Luminol chemiluminescence reaction: a new method for monitoring nitric oxide in vivo. Anal Chim Acta 458, 281–289.

12. Yao, D. C., Vlessidis, A. G., Evmiridis, N. P., Siminelakis, S., Dimitra, M. (2004) Possible mechanism for nitric oxide and oxidative stress induced pathophysiological variance in acute myocardial infarction development – a study by a flow injection-chemiluminescence method. Anal Chim Acta 505, 115–123. 13. Wang, J. N., Lu, M. Q., Yang, F. Z., Zhang, X. R., Baeyens, W. R. G., Campaña, A. M. G. (2001) Microdialysis with on-line chemiluminescence detection for the study of nitric oxide release in rat brain following traumatic injury. Anal Chim Acta 428, 173–181. 14. Miyamoto, A., Nakamura, K., Kishikawa, N., Ohba, Y., Nakashima, K., Kuroda, N. (2007) Quasi-simultaneous determination of antioxidative activities against superoxide anion and nitric oxide by a combination of sequential injection analysis and flow injection analysis with chemiluminescence detection. Anal Bioanal Chem 388, 1809–1814. 15. Hughes, M. N., Nicklin, H. G. (1968) Chemistry of pernitrites. I. Kinetics of decomposition of pernitrous acid. J Chem Soc A 2, 450–452.

Chapter 9 Aqueous Measurement of Nitric Oxide Using Membrane Inlet Mass Spectrometry David N. Silverman and Chingkuang Tu Abstract Membrane inlet mass spectrometry for the measurement of nitric oxide in aqueous solution provides a direct, continuous, and quantitative determination over long periods of time. The method uses a membrane that is permeable to nitric oxide and separates solution or cell suspension from a partial vacuum leading to the ionization source of a mass spectrometer. The construction of the probe varies depending on use; this report describes an inlet probe comprising a 1.0 cm segment of silicon rubber tubing attached to the vacuum inlet of the mass spectrometer. The probe is immersed in solution or suspension and in the system described here has a response time of 5–7 s and a lower detection limit of 0.5 nM nitric oxide. This apparatus was used to measure the generation of nitric oxide in solutions of NONOates and from the reactions of nitrite with hemoglobin. The usefulness of such an inlet in measuring nitric oxide in physiological systems is discussed. Key words: Nitric oxide, mass spectrometry, membrane inlet, nitrite, hemoglobin.

1. Introduction Membrane inlet mass spectrometry (MIMS) to measure nitric oxide (NO) in aqueous solution is based on the use of membranes permeable especially to uncharged molecules of low molecular weight, with NO measured by the m/z 30 peak. The membrane separates the solution or suspension from a partial vacuum that leads to the ionization source of a mass spectrometer. This method was first devised many decades ago by Hoch and Kok (1) who pointed out its usefulness in measuring CO2 , O2 , and N2 in studies of algal and plant physiology (1). Since that date H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_9, © Springer Science+Business Media, LLC 2011

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the method has been modified and its applications diversified in many studies. Reviews have covered MIMS applied to biological, chemical, and environmental issues (2–6). The use of MIMS is a well-described method, especially in the measurement of volatile organic compounds (4), of CO2 in physiological studies (7–9), and in understanding the mechanism of carbonic anhydrase (10). These devices have been used to detect non-polar, low molecular weight organic compounds in blood (4, 11) or following wastewater treatment (12). The application of MIMS for detecting NO in aqueous solutions as well as gas phase was introduced by Lewis et al. (2) who provided many references to the use of membrane-inlet technology for the detection of dissolved, nonpolar molecules. However, despite the advantage of a quantitative and real time, direct measure of NO in solution, MIMS has not been applied extensively to studies of NO.

2. Materials 2.1. Variety of Inlets

The construction of the inlet itself varies in different reports. In some constructions, solutions or cell suspensions to be analyzed are placed in a vessel the bottom of which is the permeable membrane supported by a porous disk (1, 13); here a paddle or magnetic stirrer is used to avoid diffusion limitations. The version described below uses as the membrane inlet probe a small (1 cm) segment of silicon rubber tubing attached to a glass tube that leads to a mass spectrometer, a version which has the advantage that it can be immersed in repetitive manner in a number of solutions or suspensions to be tested.

2.2. Semi-permeable Membrane

The material used for the permeable membrane determines the classes of permeable molecules that will pass through the membrane and be detected. Most often silicon rubber (Silastic, Dow Corning Corporation) is used because it is hydrophobic and has reduced permeability to water that lessens the entry of water vapor into the mass spectrometer (see Notes 1 and 2).

2.3. Membrane Inlet

The inlet probe is made from a length of silastic tubing (1.5 mm i.d. and 2.0 mm o.d.), which was sealed at one end by a glass bead and attached at the other end to a piece of glass tubing leading to the ion source of an Extrel EXM-200 mass spectrometer (Fig. 9.1) (see Notes 3 and 4). The silastic tubing was prevented from collapsing in the partial vacuum of the inlet by a helical coil of fine stainless steel wire, and the length of silastic tubing between the glass bead and glass tubing was 1.0 cm. The configuration of membrane inlet used here is very similar to that

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Fig. 9.1. The membrane inlet which is immersed in solution or cell suspension for mass spectrometric measurements: (1) Wire helix for support of the Silastic tubing; (2) Silastic tubing approximately 1 cm in length; (3) Glass bead to seal the Silastic tubing. Taken from (15).

of Brodbelt et al. (4) that was used to detect dichloromethane, methoxyflurane, and styrene in blood (see Notes 5 and 6). 2.4. Reaction Vessel

Any convenient vessel can be used since the membrane inlet is immersed in solution or suspension. Use a 3 mL glass cuvette (1.0 cm pathlength standard cuvette) that is modified for the introduction of samples and inert gas and is sealable by injection septums and teflon screw plugs with outflow via a second syringe (14). Place a small magnetic stir bar in the vessel (see Note 7). Bubble helium to purge extraneous gases and stop when the mass spectrometer indicates these gaseous solutes are at an acceptable level. Inject solutions containing reagents to generate NO through a syringe into the reaction vessel. Immerse the vessel itself in a water bath for temperature control. In our configuration, this airtight vessel was inserted into a spectrophotometer (HewlettPackard 8453) for concomitant absorption measurements. Simultaneous measurements by MIMS and optical measurements were performed in experiments with hemoglobin (15) and are easily done when absorption changes accompany the reactions of NO.

2.5. Spectrometer Parameters

Electron impact ionization (70 eV) was used at an emission current of 1 mA. Source pressures were approximately 1 × 10–6 torr. The resulting spectra were well resolved with a return of the ion current (detector response) to the baseline separating each mass unit.

3. Methods 3.1. Calibration

Calibrate the inlet of Fig. 9.1 and mass spectrometer by injecting solutions containing known concentrations of NO into buffered solutions in the reaction vessel (14). Two methods can be used to prepare solutions of known NO concentration. First, bubble NO gas from a gas bottle into water at 25◦ C until saturated with NO

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(16). Prepare solutions by diluting with degassed buffer using airtight syringes. Second, generate NO in solution by reduction of nitrite using 0.1 M KI in 0.1 M HCl (17). Upon rapid addition of NO into the reaction vessel, the ion current at the mass spectrometer is recorded when it reaches a maximal plateau. In our apparatus, the injections caused an immediate increase in the m/z 30 peak which was first order with a half-time of 5–7 s. A plot of maximal ion current versus NO concentration should be linear (Fig. 9.2). In our hands this linearity extended to the highest concentration of NO we measured, 20 µM. At the low concentration range, we were able to extend this linearity down to about 0.5 nM (Fig. 9.3). This limit is determined predominantly by the properties of the silicon rubber tube, its thickness and permeability to NO, and the surface area of the tube that is exposed to solution. In principle, changing these properties could enhance sensitivity.

Fig. 9.2. The ion current (arbitrary scale) at m/z 30 measured with the membrane inlet mass spectrometer using solutions containing dissolved NO of known concentrations. Solutions were prepared by addition of a stock solution of dissolved NO into buffered, degassed solutions in the reaction vessel (25 mM Mes, pH 6.9, 25◦ C). The solid line is a least squares fit in a linear regression with correlation coefficient 0.998. Data from (14).

3.2. Generation of NO from NONOates

Typical data using the membrane inlet are shown in Fig. 9.4 for the generation of NO using a NONOate. The NONOates are NO adducts that release NO into solution under designated conditions. Use MAHMA NONOate ((Z)-1-[N-methyl-N-[6-(Nmethylammoniohexyl)amino]]diazen-1-ium-1,2-diolate) (18) to generate NO in the membrane inlet apparatus and measure the

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Fig. 9.3. An extension of the ion current to lower concentrations of NO showing the lower limits of detection of the membrane inlet mass spectrometer. Conditions as in Fig. 9.2.

Fig. 9.4. Time course of the ion currents in the generation of NO from MAHMA NONOate added at the time zero at a concentration of 1.2 µM after mixing. The data are (from the top) the detector response at m/z 30, m/z 46, and m/z 76. The solution contained 50 mM phosphate buffer, 78 mM NaCl, and 1 mM EDTA at pH 6.7 and 25◦ C (inset). Time course for the generation of NO from MAHMA NONOate expressed in units of concentration (nM). The accumulation of NO was first order with a half-time of 35 s. At longer times, the half-time for decrease at m/z 30 was approximately 10 min. Taken from (14).

generation of NO. Dissolve MAHMA NONOate in 10 mM NaOH (conditions under which it is stable) and without delay dilute (about 1:1,000 v/v) into well-buffered solutions (50 mM sodium phosphate buffer, 78 mM NaCl at 25◦ C and pH 6.7) in the reaction vessel. Figure 9.4 shows the generation of NO for a typical experiment. The resulting increase in the mass peak at m/z 30 should adequately fit to a first-order increase with a

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half-life of 35 s for MAHMA NONOate. At pH 7.4 the half-life for the generation of NO was 2.0 min which was compared with a published value of 2.7 min under somewhat different conditions (22◦ C, 100 mM phosphate buffer at pH 7.4) (18) (see Note 8). In the experiment of Fig. 9.4, the decrease in the m/z 30 peak at longer times has a half-life of approximately 10 min and presumably represents the loss of NO from solution across the membrane inlet into the mass spectrometer and also into the headspace of the reaction vessel. Figure 9.4 also demonstrates that the experiment detects no increases in peaks at m/z 46 or 76 at which NO2 and N2 O3 would occur, a result that is obtained even at oxygen levels near saturation (atmospheric pressure) (see Note 9). 3.3. Accumulation of NO After Addition of Nitrite to Hemoglobin Solutions

Add sodium nitrite (8 mM) to a solution of 38 µM deoxyHb(FeII ) (heme concentration) to observe a small lag of about 3 min in which the rate of free, unbound NO accumulation in solution as detected by the m/z 30 peak is small. This is followed by an increased phase of NO accumulation starting at about 3 min (Fig. 9.5). Again, the membrane inlet method is detecting free, unbound NO in solution. In this initial phase, the concentrations of NO are less than in the control which contains no hemoglobin (Fig. 9.5). This most likely occurs because NO generated in the uncatalyzed reactions of nitrite (19) and NO generated by the interaction of nitrite with hemoglobin (20, 21) is bound avidly to deoxy-Hb(FeII ) (15). After about 3 min, the deoxyHb(FeII ) is saturated with NO and the accumulation of NO proceeds in a second phase to about 14 nM in Fig. 9.5. The mechanism of this latter phase of NO accumulation is uncertain, but is

Fig. 9.5. The time course of dissolved NO concentrations (obtained from m/z 30) upon addition of nitrite to solutions containing 38 µM deoxy-Hb(FeII ), or 38 µM oxy-Hb(FeII ), or no hemoglobin. (These are heme concentrations.) At time zero, NaNO2 was added to attain a concentration of 8 mM. Solutions also contained 50 mM phosphate buffer at pH 6.8, 110 mM NaCl, 2 mM EDTA at 23◦ C. Data taken from (15).

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perhaps due to the reaction of nitrite with met-Hb(FeIII ) or with NO-Hb(FeII ). Figure 9.5 shows that addition of nitrite at 8 mM to a solution of 38 µM oxy-Hb(FeII ) results in no detectable NO, because reactions with oxy-Hb(FeII ) have depleted NO from solution. The reaction of NO with oxy-Hb(FeII ) is complex, but the mechanism resulting in nitrite and met-Hb(FeIII ) production is discussed (22–25). These data are of physiological relevance in that they show no pulse of NO accumulation after addition of supra-physiological concentrations of nitrite. Thus the role of erythrocytes in producing vasodilatory levels of NO (>1 nM) from blood nitrite (in the order of 1 µM) is uncertain. This is just one piece of information in a much discussed topic (21, 26, 27) (see Note 10). 3.4. Overview

We show here the applicability of MIMS to the measurement of NO in solution. The use of such an inlet to measure NO has several advantages, some of which we have demonstrated in this chapter. This provides a direct, continuous, and quantitative determination of NO concentrations over long periods of time. In addition, the device allows the concomitant and quantitative determination of a number of other volatile compounds of physiological interest such as O2 and CO2 . Proposed intermediates in reactions involving NO, such as NO2 and N2 O3 , could possibly be detected if conditions could be found in which they accumulate. In addition the use of stable isotope labeling is possible.

4. Notes 1. Although silicon rubber is most often used as the permeable membrane, other materials such as Teflon (1, 13) and polyethylene (1) have been used. Quantitative expressions for the rate of transport of volatile solutes including NO across the membrane in MIMS have been presented (1, 2); this includes consideration of the area and thickness of the membrane, the solubility and diffusivity of NO in the membrane material, and the partial pressure across the membrane as the driving force for NO movement. 2. To capture water vapor that passed across the inlet, a dry ice-acetone trap was placed between the membrane inlet and the ion source; however, elimination of this trap did not affect the response time of the system to NO. 3. This inlet is simple and straightforward to construct, perhaps offsetting the disadvantage of needing a mass

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spectrometer to make these measurements. The device is easily adaptable to miniaturization or further adaptation by altering membrane material or dimensions. 4. In our initial apparatus, the distance from the Silastic inlet to the ion source was 120 cm, but this is too long and contributed to the response time of several seconds. 5. The membrane inlets are also applicable for measurement of molecules in the gas phase, as would be present, for example, in the headspace above solutions or suspensions. These properties in addition to the high selectivity and sensitivity of the method are strong advantages in the use of MIMS in detection of NO. 6. In other variations, a flow system is used in which the sample to be analyzed is passed through a small tube which is permeable to molecules of small molecular weight (2, 11, 28). Yet another rather unique version used a hypodermic needle with a 0.07 mm hole covered or filled with permeable silicon rubber (29). 7. It is important to maintain a stirring rate in solution or the NO passing across the membrane will be determined in part by diffusion to the site of the membrane inlet. 8. Studies varying the temperature from 0 to 37◦ C have not been a problem. Also a wide range of buffers and values of pH of solution in contact with the membrane have been used. 9. This membrane inlet technology would be useful in determining very fast reaction rates of NO chemistry if used in a stopped-flow or continuous-flow apparatus. For the measurement of more rapid processes, a flow cell would be appropriate in which solutions are passed through a tube of silicon rubber (2). 10. Silastic does not promote coagulation of blood and is rather resistant to caking or sticking. Hence, this membrane inlet is suitable for physiological measurements with cultured cells in vitro and in vivo with intraluminal insertion of the probe into a blood vessel similar to a catheter.

Acknowledgment We thank Dr. Erik Swenson who first suggested this project to us. Work on this research was supported by funds from the University of Florida and NIH GM25154.

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References 1. Hoch, G., Kok, B. (1963) A mass spectrometer inlet system for sampling gases dissolved in liquid phases. Arch Biochem Biophys 101, 160–170. 2. Lewis, R. S., Deen, W. M., Tannenbaum, S. R., Wishnook, J. S. (1993) Membrane mass spectrometer inlet for quantitation of nitric oxide. Biol Mass Spectrom 22, 45–52. 3. Lauritsen, F. R., Lloyd, D. (1994) Directdetection of volatile metabolites produced by microorganisms – membrane inlet massspectrometry. Mass Spectrom Charact Micro 541, 91–106. 4. Brodbelt, J. S., Cooks, R. G., Tou, J. C., Kallos, G. J., Dryzga, M. D. (1987) In vivo mass-spectrometric determination of organic-compounds in blood with a membrane probe. Anal Chem 59, 454–458. 5. Kotiaho, T., Lauritsen, F. R., Choudhury, T. K., Cooks, R. G., Tsao, G. T. (1991) Membrane introduction mass-spectrometry. Anal Chem 63, 875–886. 6. Lauritsen, F. R., Kotiaho, T., Choudhury, T. K., Cooks, R. G. (1992) Direct detection and identification of volatile organic-compounds dissolved in organic-solvents by reversedphase membrane introduction tandem massspectrometry. Anal Chem 64, 1205–1211. 7. Tu, C., Wynns, G. C., McMurray, R. E., Silverman, D. N. (1978) CO2 kinetics in redcell suspensions measured by O-18 exchange. J Biol Chem 253, 8178–8184. 8. Itada, N., Forster, R. E. (1977) Carbonicanhydrase activity in intact red blood-cells measured with O-18 exchange. J Biol Chem 252, 3881–3890. 9. Gerster, R. (1971) Kinetics of oxygen exchange between gaseous C18O2 and water. Int J Appl Radiat Isot 22, 339–348. 10. Silverman, D. N. (1982) Carbonic anhydrase: oxygen-18 exchange catalyzed by an enzyme with rate-contributing protontransfer steps. Methods Enzymol 87, 732–752. 11. Trushina, E. V., Clarke, N. J., Benson, L. M., Tomlinson, A. J., McMurray, C. T., Naylor, S. (1998) A miniaturized membrane inlet mass spectrometry interface for analysis of nitric oxide in human plasma. Rapid Commun Mass Spectrom 12, 985–987. 12. Calvo, K. C., Weisenberger, C. R., Anderson, L. B., Klapper, M. H. (1981) Permeable membrane – mass-spectrometric measurement of reaction-kinetics. Anal Chem 53, 981–985. 13. Conrath, U., Amoroso, G., Kohle, H., Sultemeyer, D. F. (2004) Non-invasive online detection of nitric oxide from plants and

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some other organisms by mass spectrometry. Plant J 38, 1015–1022. Tu, C. K., Swenson, E. R., Silverman, D. N. (2007) Membrane inlet for mass spectrometric measurement of nitric oxide. Free Radic Biol Med 43, 1453–1457. Tu, C. K., Mikulski, R., Swenson, E. R., Silverman, D. N. (2009) Reactions of nitrite with hemoglobin measured by membrane inlet mass spectrometry. Free Radic Biol Med 46, 14–19. Lide, D. R. (1998) CRC Handbook of Chemistry and Physics. CRC-Press, Boca Raton, FL. Garside, C. (1982) A chemi-luminescent technique for the determination of nanomolar concentrations of nitrate and nitrite in seawater. Marine Chem 11, 159–167. Hrabie, J. A., Klose, J. R., Wink, D. A., Keefer, L. K. (1993) New nitric oxide-releasing zwitterions derived from polyamines. J Org Chem 58, 1472–1476. Samouilov, A., Kuppusamy, P., Zweier, J. L. (1998) Evaluation of the magnitude and rate of nitric oxide production from nitrite in biological systems. Arch Biochem Biophys 357, 1–7. Huang, K., Keszler, A., Patel, N., Patel, R., Gladwin, M., Kim-Shapiro, D., et al. (2005) The reaction between nitrite and deoxyhemoglobin: reassessment of reaction kinetics and stoichiometry. J Biol Chem 35, 31126–31131. Singel, D. J., Stamler, J. S. (2005) Chemical physiology of blood flow regulation by red blood cells: the role of nitric oxide and S-nitrosohemoglobin. Annu Rev Physiol 67, 99–145. Doyle, M. P., Pickering, R. A., DeWeert, T. M., Hoekstra, J. W., Pater, D. (1981) Kinetics and mechanism of the oxidation of human deoxyhemoglobin by nitrites. J Biol Chem 256, 12393–12398. Doyle, M. P., Hoekstra, J. W. (1981) Oxidation of nitrogen-oxides by bound dioxygen in hemoproteins. J Inorg Biochem 14, 351–358. Herold, S., Exner, M., Nauser, T. (2001) Kinetic and mechanistic studies of the NO center dot-mediated oxidation of oxymyoglobin and oxyhemoglobin. Biochemistry 40, 3385–3395. Herold, S., Rock, G. (2005) Mechanistic studies of the oxygen-mediated oxidation of nitrosylhemoglobin. Biochemistry 44, 6223–6231. Basu, S., Grubina, R., Huang, J., Conradie, J., Huang, Z., Jeffers, A., et al.

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(2007) Catalytic generation of N2 O3 by the concerted nitrite reductase and anhydrase activity of hemoglobin. Nat Chem Biol 3, 785–794. 27. Li, H. T., Cui, H. M., Kundu, T. K., Alzawahra, W., Zweier, J. L. (2008) Nitric oxide production from nitrite occurs primarily in tissues not in the blood – critical role of xanthine oxidase and aldehyde oxidase. J Biol Chem 283, 17855–17863. 28. Southan, G. J., Srinivasan, A. (1998) Nitrogen oxides and hydroxyguanidines:

formation of donors of nitric and nitrous oxides and possible relevance to nitrous oxide formation by nitric oxide synthase. Nitric Oxide Biol Ch 2, 270–286. 29. Lloyd, D., Thomas, K., Price, D., ONeil, B., Oliver, K., Williams, T. N. (1996) A membrane-inlet mass spectrometer miniprobe for the direct simultaneous measurement of multiple gas species with spatial resolution of 1 mm. J Microbiol Methods 25, 145–151.

Chapter 10 Quantum Cascade Laser Technology for the Ultrasensitive Detection of Low-Level Nitric Oxide Angela Elia, Pietro Mario Lugarà, Cinzia Di Franco, and Vincenzo Spagnolo Abstract Several spectroscopic methods based on mid-infrared quantum cascade lasers for the ultrasensitive detection of nitric oxide have been developed with detection limit in ppbv and sub-ppbv range. We will describe here a selection of the most effective techniques, i.e., laser absorption spectroscopy, cavityenhanced spectroscopy, photoacoustic spectroscopy, and Faraday modulation spectroscopy. For each technique, advantages and drawbacks will be underlined. Key words: Nitric oxide detection, quantum cascade lasers, absorption spectroscopy, cavityenhanced spectroscopy, photoacoustic spectroscopy, Faraday modulation spectroscopy.

1. Introduction The development of compact optical sensors for nitric oxide (NO) detection is of interest for different applications, such as environmental monitoring (1), vehicle exhaust control (2), industrial process control (3), and medical diagnostics (4). Both optical and nonoptical analytical methods have been developed to measure ultralow concentrations of NO. Nonoptical approaches include mass spectrometry and gas chromatography. The main drawbacks of these techniques are size and cost of the apparatus, the need for sample conditioning, consumables, and the inability to make real-time and online measurements. The most advanced optical techniques are based upon either chemiluminescence or laser absorption processes. In particular, infrared H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_10, © Springer Science+Business Media, LLC 2011

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laser absorption spectroscopy (LAS) has become an extremely effective tool for the detection and quantification of molecular trace gases. The detection sensitivity of LAS ranges from ppmv (part per million in volume) down to pptv (part per trillion in volume) levels depending on the specific gas species and the detection method employed. Several infrared spectroscopy techniques have been developed for NO monitoring in ppbv (part per billion in volume) and subppbv range. NO is a polar molecule and infrared (IR) active. The gasphase NO molecule has two absorption bands near 5.2 and 2.6 µm. For highly sensitive spectroscopic trace detection, a suitable band must be selected. It should be characterized by strong absorption intensity, but isolated from other interfering species, i.e., water and carbon dioxide. In particular, the detection of NO is more effective in the mid-IR spectral region around 5.2 µm, where the strongest spectral feature has intensity lines of about ∼6.04 × 10–20 cm/molecule and is separated from interference absorption bands. Several types of laser sources are available in this spectral region. These include line-tunable carbon monoxide (CO) lasers, lead–salt diode lasers, quantum cascade lasers, and nonlinear laser sources such as optical parametric oscillators (OPO) and difference frequency generation (DFG) systems. The ideal source for spectroscopic applications should have the following characteristics: (i) high optical power, to get high laser signal-to-noise ratios; (ii) narrow line width, to obtain good selectivity; (iii) single mode operation; (iv) low source noise and low amplitude fluctuations; (v) high stability to environmental conditions, i.e., temperature, pressure, humidity, and vibrations; (vi) high reliability; and (vii) compact and robust overall sensor package size. The lead–salt lasers are difficult to incorporate into a commercial device because of their need for cryogenic cooling. Nonlinear generation of IR light via DFG or OPO (based on periodically poled lithium niobate crystals) provides a broad continuous tuning range (hundreds of cm–1 ) (5–8). However, to reach the ppbv level of sensitivity with the low power achievable by DFG (up to few mW) (9–12), advanced detection schemes are needed. OPOs have relatively high power levels (1 W, continuous wave operation) and narrow line width (typically a few MHz over 1 s) and, therefore, represent an excellent source for sensitive spectroscopic gas analysis. In combination with fiber pump laser technology, the OPO-based sources offer the attractive advantage of a rather compact setup. However, to date, OPOs are less suitable for field applications as the cavity needs occasional tweaking. Until a few years ago, direct generation of tunable mid-IR radiation using solid-state lasers suffered especially from limited output

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power, low-temperature operation, and limited tuning properties. Instead, quantum cascade lasers (QCLs) represent a valid choice, since they overcome some of the major drawbacks of other traditional mid-IR laser sources. These include the lack of continuous wavelength tunability, the large size and weight of gas lasers, low output power and cooling requirement of lead–salt diode lasers, and the complexity of nonlinear optical generators. Several very effective approaches utilizing QCLs for the optical sensing of NO have been reported.

2. Properties of Quantum Cascade Lasers

Quantum cascade lasers have been constantly developed since their invention at Bell Laboratories in 1994 (13) and so far represent the most interesting source for optical sensors. QCLs are unipolar semiconductor lasers based on intersubband transitions in a multiple quantum-well heterostructure (see Note 1). Typical emission wavelengths can be varied in the range of 3–17 µm. The innovations in QCLs led to the first demonstration of continuous wave (cw) operation at room temperature at the wavelength of 9 µm in 2002 (14). In 2003, Yu and colleagues (15) achieved room-temperature cw operation at shorter wavelengths and very large output powers by reducing the doping in the active region and ridge width. To achieve the single frequency required by chemical sensing applications, a Bragg grating was integrated into the laser waveguide for the first time at Bell Laboratories by Gmachl and coworkers, resulting in a distributed-feedback (DFB) laser operating at cryogenic temperatures (16). The latest generation of QC-DFB lasers is based on a “topgrating” approach that takes advantage of the characteristics of a mid-IR waveguide. For mid-IR wavelengths below 15 µm, dielectric waveguides of low-doped semiconductor layers with a proper refractive index modulation are used (17). Furthermore, roomtemperature commercial cw DFB-QCLs with an optical power larger than 100 mW (18, 19) and prototype lasers emitting up to few watts (20) have been developed. QCLs generally require several amperes of current in cw operation, and compliance voltages of 5–10 V. The resulting thermal load to the laser is substantial, where good thermal management is necessary to reach room-temperature operation. In addition, the tuning range of a DFB-QCL covers one or two absorption lines of a gas. However, some applications are based on the detection of multicomponent gas matrix, requiring a large tuning range. Fortunately, the inter-subband transitions can be tailored to enable

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the design of active regions with very large gain bandwidth. In 2004, Maulini and coworkers (21) demonstrated the first external cavity QCL (EC-QCL) operating in a cw over a record frequency span of 175/cm, using a bound-to-continuum QC structure with an optical power up to 10 mW. Coarse tuning can be obtained by rotating the grating, while changing the cavity length and laser chip temperature allows the fine tuning. The main advantage of EC-QCLs with respect to DFB-QCLs is a broader tuning range, limited only by the spectral bandwidth of its gain element. A broader tunability of several hundreds of wave numbers will allow the detection of entire absorption bands and enhance the flexibility of QCLs for trace gas analysis. The usefulness of these lasers for spectroscopic applications has recently been demonstrated by Wysocki et al. (3) who used a thermoelectrically (TE) cooled cw EC-QCL for spectroscopic absorption measurement of NO (22).

3. Methods 3.1. Absorption Spectroscopy

Laser absorption spectroscopy (LAS) has a great potential for the detection and monitoring of trace gases. It operates on the principle that the amount of light absorbed by a gas is related to the concentration of the target species (see Note 2). For each gas a strong absorption line, preferably free of interference due to other gases in the sample cell, must be selected. The strongest molecular rotational–vibrational transitions, which are desired to perform ultrasensitive concentration measurement, are in the mid-IR spectral region (see Note 3). The most important advantage of LAS is the ability to provide absolute quantitative assessments of species. Its biggest disadvantage relies on the measurement of a small change in a high level optical power; any noise introduced by the light source or the transmission through the optical system will decrease the sensitivity of the technique. Laser absorption spectrometric techniques are therefore often limited to detection of absorbance ∼10–3 , which is far away from the theoretical shot noise level, which for a single-pass technique is in the 10–7 –10–8 range. This sensitivity is insufficient for many types of applications. Obtaining detection sensitivities at ppbv or sub-ppbv levels requires either long effective optical path lengths or suppression of laser and optical noise. Long optical path lengths are typically obtained in multipass absorption cells. There are three types of multipass cells in use (23–26): White cells (see Note 4), Herriott cells (see Note 5), and astigmatic mirror cells (see Note 6).

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The designs of the three types of cells must fulfill specific rules so that the beam goes out of the cell after a controlled number of passes, especially in the case of astigmatic cells. Optical output of the cell decreases exponentially with the number of passes. In order to have small absorptions and maximum sensitivity, the optimal number of reflections corresponds to the output light being 1/e times the input light. However, as long as the noise is determined by laser intensity fluctuations, rather than by detector noise, additional passes will improve the signal-to-noise ratio (SNR). In 2006, Moeskops and coworkers (27) monitored the unresolved NO doublet at 1,850.18/cm with a detection limit of 0.2 ppbv corresponding to a minimal detectable absorption of 8.8 × 10–9 /cm Hz1/2 (see Note 7). Atmospheric NO was detected by MacManus and coworkers (28) via a totally noncryogenic spectrometer with a detection limit of 0.1 ppbv in pulsed mode and 0.03 ppbv in cw after an average time of 30 s (see Note 8). The use of a single-pass cell resulted in an absorption spectrometer less sensitive, as reported by Kasyutich and coworkers (29). They reported a minimum detection limit of 2.7 ppmv in plasma diagnostics (see Note 9). 3.2. Cavity-Enhanced Spectroscopy

Cavity-enhanced spectroscopy (CES) methods provide a much higher sensitivity than conventional long optical path length absorption spectroscopy (see Note 10). Different techniques have been implemented. In particular, cavity ring-down spectroscopy (CRDS) is the most popular CES embodiment. It is a direct absorption technique which can be performed with pulsed or continuous light sources. It is based on the measurement of the decay time of an injected laser beam in a high-finesse optical cavity in the presence of an absorbing gas by measuring the time dependence of the light leaking out of the cavity (see Note 11). The technique based on cw laser sources was first proposed by Romanini et al. (30) using cw near-IR DFB diode lasers and was extended to QCL sources by Paldus et al. (31). CRDS with cw QC-DFB laser has been successfully applied to the detection of NO at ppbv concentrations by Kosterev et al. (see Note 12) (32). The schematic NO sensor is shown in Fig. 10.1 (see Note 13). The authors demonstrated the detection of NO in pure N2 with concentrations at ppbv levels with a 0.7-ppbv standard error for a data collection time of 8 s and with 4,000 ring-down events. It was not possible to use this sensor directly for measurements of NO concentration in exhaled air (10 ppbv) because of a strong interference with CO. Another technique based on a high-finesse optical cavity is called “integrated cavity output spectroscopy” (ICOS) or

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Fig. 10.1. Schematic of a CRDS-based gas sensor. r1 : current monitor resistor; r2 : current-limiting resistor. Two wedged ZnSe windows shown near the reference cell were used to form an etalon for fine frequency calibration. (Reproduced from ref. (32) with kind permission of The Optical Society of America.)

“cavity-enhanced absorption spectroscopy” (CEAS). Laser light is coupled into the high-finesse cavity via accidental coincidences of the laser frequencies and the cavity eigenmodes. The alignment of the cavity mirrors and the laser beam are optimized in order to maximize the number of transverse modes excited. Intensity radiation leaking out of the optical cavity is time-integrated and averaged over many cavity modes. Subsequently, the intensity radiation is inversely proportional to the total cavity losses which can be used to determine the absorption of the intracavity medium (see Note 14). This approach was demonstrated with a cw QC-DFB laser in 2001 (33) for the detection of NO in human breath for biomedical applications (see Note 15). The detection limit was found to be 16 ppbv and was limited by a baseline noise of 1% (averaging 10 QC laser scans) which is intrinsic to this technique and results from the mode structure of the cavity transmission spectrum due to the incomplete averaging of cavity modes. Some improvements in the baseline noise can be achieved with a recently developed off-axis ICOS (OA-ICOS). In this configuration, the laser beam is directed off-axis with respect to the cavity axis in order to increase the spectral density of cavity modes and thus minimize the noise in the absorption spectra by improving the averaging of the cavity output. The off-axis ICOS measurement technique requires a less critical alignment of the exciting laser beam and is more insensitive to vibrations and misalignments than CRDS and on-axis ICOS.

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Several QCL-based off-axis ICOS systems for the detection of NO have been reported with sensitivities in the ppbv level (34–38). The first NO analyzer based on an off-axis ICOS and cw liquid-N2 -cooled QCL was developed in 2004 to measure NO concentrations in exhaled human breath (34). In combination with a wavelength modulation technique, a noise-equivalent (SNR = 1) sensitivity of 2 ppbv in a nasal breath sample was demonstrated for a 15 s data acquisition and integration time. Two years later, an off-axis ICOS sensor (36, 37) based on a thermoelectrically cooled DFB-QCL operating in cw mode at 5.45 µm combined with a wavelength modulation technique was developed to measure NO concentrations at the sub-ppbv levels. A schematic of the sensor is reported in Fig. 10.2. A noise-equivalent minimum detection limit of 0.7 ppbv with a 1 s observation time was achieved (36) in N2 and 1.2 ppbv with a 4 s observation time in exhaled breath samples (37).

Fig. 10.2. TEC-cw-DFB-QCL-based OA-ICOS sensor. MCT is a cryogenically cooled photovoltaic HgCdTe detector and MCZT is a thermoelectrically cooled HgCdZnTe photodetector. (Reproduced from ref. 36 with kind permission of Springer Science and Business Media.)

More recently, McCurdy et al. (38) have reported an off-axis ICOS sensor capable of real-time detection of NO and CO2 in a single breath cycle, achieving a NO detection limit of 0.4 ppbv with a 1 s integration time, in good agreement with the data acquired with a commercial chemiluminescence NO gas analyzer. 3.3. Photoacoustic Spectroscopy

Photoacoustic spectroscopy (PAS) represents an effective method for sensitive trace gas detection. PAS is an indirect technique in

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which the effect on the absorbing medium and not the direct light attenuation is detected. In particular, it is based on the generation of an acoustic wave resulting from the absorption of modulated light of appropriate wavelength by molecules. The amplitude of this sound wave is directly proportional to the gas concentration and can be detected with a sensitive microphone if the laser beam is modulated in the audio frequency range (see Note 16). In combination with QCLs, PAS offers the advantage of high sensitivity (ppbv detection limits), large dynamic range, compact setup, fast time-response, and simple optical alignment, if compared with other competing detection schemes, such as multipass absorption spectroscopy or cavity ring-down spectroscopy, which offer similar performances but require more sophisticated equipments. PAS with DFB-QC laser for the detection of NO has been successfully demonstrated in 2005 by Elia et al. (39); a detection limit of 500 ppbv has been reported. The photoacoustic sensor for the detection of NO consisted of a commercially available distributed feedback quantum cascade laser source, a resonant photoacoustic cell, and a signal acquisition and processing equipment (see Note 17). In Fig. 10.3, a schematic diagram of the optoacoustic sensor is reported.

Fig. 10.3. Schematic diagram of the optoacoustic sensor.

More recently, the same group obtained a detection limit of 150 ppbv with a 10 s integration time constant for the detection of NO (40). This improved result is mostly due to improvements in the PA cell and damping of electromagnetic noise sources. The Groupe de Spectrometrie Moleculaire et Atmospherique (Reims, France) developed a Helmholtz resonant photoacoustic sensor for NO detection (see Note 18). An extrapolated detection limit of 20 ppb of NO in nitrogen with a laser power of 3 mW was demonstrated (41). The presence in this region of interferences from water vapor lines prevents NO detection in air with a detection limit lower than 1 ppmv. More recently Spagnolo et al. (42) demonstrated a NO sensor based on quartz-enhanced photoacoustic (QEPAS) detection and an external cavity (EC) quantum cascade laser. The key innovation of QEPAS is to detect optically generated sound using a sharply resonant piezoelectric quartz tuning fork transducer. The NO concentration resulting in

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a noise-equivalent signal was found to be 15 ppbv, with 100-mW optical excitation power and a data acquisition time of 5 s.

3.4. Faraday Spectroscopy

Faraday modulation spectroscopy (FMS) is an alternative detection technique capable of enhancing the sensitivity of LAS for radicals and ions (43–45). The absorption lines of radicals can be tuned by an external magnetic field that breaks the magnetic degeneracy of the rotational states (Zeeman effect). The resulting frequency shift of transitions is different for left-handed and righthanded circularly polarized light, giving rise to different refractive indices for these polarization components at a given radiation wavelength (circular birefringence). As a light beam, originally linearly polarized, propagates through the sample, this anisotropy leads to a rotation of the polarization axis. This magnetically induced birefringence in a longitudinal field and the related rotation of the polarization axis of linearly polarized light is called Faraday effect. The rotation is detected by means of putting the sample between nearly crossed polarizers. In this way, laser amplitude noise is largely suppressed. Also, employing a static magnetic field B > 0 in combination with a tunable laser, the sensitivity of direct absorption spectroscopy can be improved by 2–3 orders of magnitude. Since NO is the only radical in ambient air with a spectrum near λ = 5.2 µm, this spectroscopic approach coupled with QCLs is advantageous since there is no interference from other constituents in the air sample. Also, this technique enables the detection of NO at low ppbv concentration levels that are typical for biomedical applications. Thus, FMS combined with polarization detection is one of the most sensitive spectroscopic methods in the mid-IR wavelength range. The typical experimental setup of FMS employing QCLs is shown in Fig. 10.4 (43). The QCL source emits in the wavelength region around 5.2 µm. The laser beam passes through a Rochon polarizer and is then fed through a detection cell which is inside a copper wire coil for the application of a magnetic field. A second Rochon polarizer behind the cell is set to the crossed position with a slight offset angle. The transmitted portion is focused by means of a parabolic mirror to a liquid-N2 -cooled indiumantimonide (InSb) photodetector. If the QCL frequency is in resonance with a NO transition, and a magnetic field is applied, the polarization axis of the laser beam is slightly rotated due to the Faraday effect. Thus, a corresponding part of the light passes through the analyzer and reaches the detector (see Note 19). The obtained signal is proportional to the NO concentration inside the cell. Since a continuously tunable QCL is used, a servo loop is needed to stabilize the laser frequency to the NO absorption frequency. The setup of the frequency stabilization is also shown in Fig. 10.4.

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Fig. 10.4. Schematic of the Faraday modulation spectrometer. Both the spectrometer and the laser frequency stabilization utilize the fact that NO is a radical and, thus, transition frequencies are tunable with a magnetic field. For sensitive detection (upper part), the Faraday rotation of the polarization axis is utilized. For the frequency stabilization (lower part), the Zeeman detuning of the transition frequency is exploited. RP: Rochon polarizer; MC: magnetic coil; SG: sinus generator; PD: photodetector; M: mirror; BS: beam splitter; L: lens. (Reproduced from ref. 43 with kind permission of Springer Science+Business Media.)

A λ/4-plate transforms the linear polarization into a circular one. The circular polarized laser light passes through a cell filled with pure NO at a pressure 1 mW) because of reduced transmission through a high-finesse optical cavity and a very sensitive detector (usually cryogenically cooled) characterized by low noise. Considering that the QCLs optical power has been steadily improved in the past few years and now commercial sources with optical power up to 1 W and prototype laser emitting up to few watts are available, the detection sensitivities of these techniques will largely improve allowing use of less sensitive but thermoelectrically cooled detectors, which are more suitable for the development of portable gas sensors. 11. CRDS has a significantly higher sensitivity than conventional absorption spectroscopy thanks to the large effective path lengths (several kilometers) that can be realized in a high-finesse optical cavity (with reflectivities of R > 99.99%) with a small sample volume, and the intrinsic insensitivity to light source intensity fluctuations. 12. The laser frequency is slowly scanned across the selected absorption line of NO and one of the cavity mirrors is dithered back and forth to ensure periodic, random

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coincidences of the laser frequency with a cavity mode. When the resonance occurs and the cavity is filled with the gas sample, the laser beam is abruptly interrupted or set off-resonance, and the relaxation time of the light leaking out of the cavity is measured. The ringdown time for a two-mirror cavity with the same reflectivity R≈1 is defined by: τ=

1 l c α l + (1 − R)

where l is the cavity length, c is the speed of light, R is the reflectivity of the cavity mirrors, and α is the absorption coefficient of the sample filling the cavity. Thus, the absorption coefficient can be determined by measuring the decay rate by the following equation: 1 α= c



1 1 − τ τempty



where τ empty is the decay constant of the cavity in the presence of a nonabsorbing sample. 13. The 37-cm-long high-finesse optical cavity was formed by two concave mirrors with a 6 m radius of curvature. A cw liquid-nitrogen-cooled DFB-QCL operating at 5.2 µm was used as a tunable single-frequency light source to access the unresolved R(13.5) components of the fundamental absorption band of NO located at 1,921.599 and 1,921.601/cm. The laser current was manipulated both for laser frequency tuning and abrupt interruptions of the laser radiation. A liquid-N2 -cooled photovoltaic HgCdTe detector was used to monitor the radiation. 14. The transmission of the cavity in the case of perfect spatial coupling is given by: I = I0

(1 − R)2 2 [(1 − R) + αL]

where I0 is the initial laser power, α is the absorption coefficient, R is the reflectivity of the mirrors, and L is the cavity length. In the case of weak absorption (αL0.1 mA) nerve stimulation is applied. However, chronic systemic administration of a number of NOS inhibitors such as Nω -nitro-L-arginine methyl ester or the relatively specific eNOS inhibitor L-N(5)-(l-iminoethyl)ornithine accelerated the

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Fig. 15.3. eNOS activity in NTS contributes to baroreceptor reflex sensitivity and chronic inhibition of eNOS reduced blood pressure in SHR. a: Sites of viral transfection identified by eGFP fluorescence documented on schematized transverse sections of the dorsal medulla after Paxinos and Watson (30). b: The effect of AVV-CMV-TeNOS in the NTS of a SHR on arterial blood pressure (BP), heart rate (HR) and spontaneous baroreceptor reflex gain (sBRG). Disabling eNOS activity in NTS lowered BP and HR but increased the cardiac sBRG in the SHR. Thus, endogenous eNOS activity in the NTS plays a major role in determining the set point of arterial pressure in the SHR and contributes to maintaining high arterial blood pressure in this animal model of human hypertension. c: Effects of chronic inhibition of eNOS activity in the NTS on mean blood pressure (MBP), HR and sBRG in conscious normotensive rats. A significant increase in sBRG was observed 14, 21 and 28 days after AW-CMV-TeNOS transfection. In contrast, sBRG did not change in eGFP-transfected and saline-treated groups. In the TeNOS-transfected group, significant decreases in HR were also observed 21 and 28 days post-adenoviral injection. + p < 0.05, ++ p < 0.01 and +++ p < 0.001 values compared before and after gene transfer. ∗ p < 0.05 and ∗∗ p < 0.01 values compared to eGFP transfected group. # p < 0.05 values compared to saline-treated group.

functional recovery of neuromuscular transmission and resulted in a measurable CMAP 7 days post-injury (20). To further reveal the role of eNOS, AVV-CMV-TeNOS was used. A single intraneural injection of either AVV-CMV-eGFP or AVV-CMV-TeNOS was performed on the day of crushing. eNOS inhibition by AVVCMV-TeNOS advanced muscle reinnervation such that CMAP was evidenced in the genioglossus muscle 1 week after the nerve crushing. This was not found to be the case in AVV-CMV-eGFPinjected control animals (20) (Fig. 15.4). These observations demonstrated that inhibition of NO synthesis of endothelial origin is beneficial for motor functional recovery after nerve crush injury.

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Fig. 15.4. Chronic intraneural inhibition of eNOS using an adenoviral vector accelerates neuromuscular function recovery and axonal regeneration. I: Experimental design. a: To evaluate the time course of the neuromuscular function recovery, the CMAP, evoked by electrical stimulation (St.) of the XIIth nerve, was recorded using electrodes implanted in the genioglossus muscle. Recordings were performed in control animals and at different time points after nerve crushing. b: Axonal regeneration was tested quantifying the number of motoneurones retrogradely labelled with Fluoro-Gold (FG) after nerve crushing 10 mm proximal to the bifurcation. FG was applied to the stump at different time points post-lesion. After FG application the animals were kept alive for 7 days to allow for the retrograde transport of the marker. Inset: Highmagnification photomicrograph of FG-labelled HMNs. Scale bar = 50 µm. II a–c: CMAPs evoked in the genioglossus muscle by single shock stimulation (arrowheads point to the stimulus artefact) of XIIth nerve 7 (a), 22 (b) or 62 (c) days after intraneural injection of Ad-CMV-eGFP or Ad-CMV-TeNOS. For comparison, recordings obtained by stimulation of the left (intact) and right (crushed) XIIth nerve are illustrated. Each trace represents an average of ten individual responses. d: The ratio between the area of CMAP evoked by electrical stimulation of the right versus the left nerve at different time points. Horizontal grey bar represents the mean (S.E.M.) in control rats. e: Differences in the latency of the CMAPs evoked by stimulation of the right and the left nerves. n = 3–4 animals per experimental group. f, g: Photomicrographs of the coronal sections of the right HN showing FG-labelled motoneurones in animals injected with Ad-CMV-eGFP (f) or AdCMV-TeNOS (g) on the crushing day. FG was applied on day 2 post-crushing and the animals were perfused 7 days after FG application. Scale bar = 100 µm. h: Numbers of FG-labelled motoneurones identified in Ad-CMV-eGFP or Ad-CMVTeNOS-treated animals. n = 4 and 7 animals for Ad-CMV-eGFP and Ad-CMV-TeNOS-transduced groups, respectively. §,#,∗ p < 0.05; non-parametric Mann–Whitney U test, with respect to the control, control and Ad-CMV-eGFP-treated or Ad-CMV-TeNOS-treated groups, respectively.

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Over the past 10 years, RNAi has been developed into a powerful tool to induce loss-of-function phenotypes. Following the design of Stegmeier et al. (21), we have recently developed a miR30-based, tetracycline (Tet)-controllable and neuronal-specific lentiviral (LVV) RNA interference system targeting nNOS. To activate this system in neurones, a second LVV expressing the Tet-off transactivator tTA using bidirectional transcriptionally enhanced human synapsin 1 (SYN) promoter was co-applied (22). This yielded in a binary system (LVV-TretightGFP-miR30-shRNA/nNOS + LVV-mCMV/SYN-tTA) which has both neuronal specificity and doxycycline (Dox)-mediated temporal control. Schematic representation of the two LVV vectors is shown in Fig. 15.2b. Protocols for lentiviral preparation are described in Section 3.4. This lentiviral binary RNA interference system has also been tested to assess the role of nNOS in central cardiovascular control at the level of NTS and in injury-induced plasticity in hypoglossal motor neurones triggered by peripheral neuropathy. Such examples are described below. We observed a 69 and 55% Dox-sensitive knock-down of nNOS expression in PC12 cells (a neurone-derived rat pheochromocytoma cell line) and in the dorsal vagal complex in rats, respectively (Fig. 15.5a). By injecting the binary RNA interference system into NTS of SHR rats, we found that systolic blood pressure was significantly increased 4 weeks later (Fig. 15.5b). This is the first description of the chronic cardiovascular role of endogenous nNOS in NTS for arterial pressure control. Our data supports a role for nNOS in long-term blood pressure control in hypertensive animals where it appears to restrain further increases perhaps providing an important protective role against stroke. The inspiratory activity of XIIth nerve motoneurones is modulated by chemoreceptor input to the respiratory network and their activity can be directly correlated with end tidal CO2 (ET CO2 ) as an indirect measure of arterial blood CO2 . One week after XIIth nerve crushing the CO2 -mediated modulation of hypoglossal motoneurone (HMN) activity was depressed. Pharmacological data suggested a role for nNOS in this process (23). LVV-Tretight-GFP-miR30-shRNA/nNOS together with LVV-mCMV/SYN-tTA were locally injected into the hypoglossal nucleus 3 days before XIIth nerve crush to allow for the expression of the nNOS shRNA hairpin. This prevented the loss of functional synaptic modulation of HMNs by the CO2 -induced changes in the respiratory drive, a characteristic outcome of the axonal injury (Fig. 15.6).

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Fig. 15.5. Western blot analyses of the knock-down functions of LVV-mediated miR30-shRNA/nNOS (a) and its effect in NTS on blood pressure in SHR (b). a (1): Dox-sensitive knock-down of nNOS expression in PC12 cells (∼69% knockdown). The ratio among different viruses in each group is 1:1:1. The total viral MOI for each well is 5. – Dox, cells were cultured in the continuous absence of Dox; + Dox, cells were cultured in the continuous presence of Dox. Adenoviral vector AVV-CMV-nNOS was used to induce high-level nNOS expression in both PC12 cells since there is no endogenous nNOS expression in this cell line. LVV-SYN-WPRE served as a control vector to balance the total input of viral particles into cells. 1: AVV-CMV-nNOS + LVV-Tretight-GFP-miR30-shRNA/nNOS + LVV-SYN-WPRE; 2: AD-CMV-nNOS + LV-Tretight-GFP-miR30-shRNA/nNOS + LVV-mCMV/SYN-tTA; 3: AD-CMV-nNOS + LV-Tretight-GFP-miR30-shRNA/nNOS + LVV-mCMV/SYN-tTA + Dox; 4: AVV-CMV-nNOS + LVV-Tretight-GFP-miR30-shRNA/Luc + LVV-mCMV/SYN-tTA (negative control); 5: Baseline control without any transfection. Please note that anti-luciferase construct, LVV-Tretight-GFP-miR30shRNA/Luc, was without effect in either cell line, indicating that the nNOS knock-down was sequence-specific. a (2): Dox-sensitive knock-down of nNOS expression of DVC in rats (∼55% knock-down). The ratio among different LVVs in each group is 1:4 and the total dose was 6 × 106 infections units (ifu) per rat. Rats in groups 2, 4 were not treated with Dox. Rats in group 3 were administered Dox in drinking water post-injection for 7 days. Samples from three rats for each group were pooled together for Western blot analyses. 1: Baseline control without any viral transfection; 2: LVV-TretightGFP-miR30-shRNA/nNOS + LVV-mCMV/SYN-tTA; 3: LVV-Tretight-GFP-miR30-shRNA/nNOS + LVV-mCMV/SYN-tTA + Dox; 4: LVV-Tretight-GFP-miR30-shRNA/Luc + LVV-mCMV/SYN-tTA (negative control). b: Four weeks after injection of the LVV binary system into the NTS, arterial pressure increased in SHR (∗ p < 0.05).

2. Materials 2.1. Adenoviral Preparation

1. Human embryonic kidney (HEK) 293 cell line. 2. Dulbecco’s Modified Eagle’s Medium (DMEM) full medium supplemented with 10% fetal bovine serum (FBS), 50 U/mL penicillin and 50 µg/mL streptomycin. 3. Trypsin (0.5 g/L) and ethylenediamine tetraacetic acid (EDTA) (0.2 g/L). 4. Tube sorvall polyallomer 18.5 mL tubes. 5. Tris–HCl buffer: 0.1 M at pH 8.0; 10 mM at pH 7.5. 6. Add sufficient CsCl to 0.1 M Tris–HCl, pH 8.0, to saturate the buffer at room temperature. Store at 4◦ C. 7. PD-10 columns: pre-packed, disposable columns containing Sephadex G-25 M for rapid desalting and buffer exchange (GE-healthcare, 17-0851-01).

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Fig. 15.6. miR30-shRNA/nNOS derived from the lentiviral binary RNA interference system prevents reduction of the chemoreceptor-modulated inspiratory activity of HMN induced by XIIth nerve crushing. a: Illustrative histograms of the instantaneous firing rate (FR; in spikes/s) of HMNs after adjusting end tidal CO2 concentrations (ETCO2 , in percent) at the indicated values. Motoneurones were recorded under four different conditions: (1) control condition without crushing; (2) 7 days after XIIth nerve crushing without virus treatment; (3) 7 days after XIIth nerve crushing, animals received microinjection in the hypoglossal nucleus of the binary RNA interference system 3 days before crushing; and (4) same as (3) except that animals are administered with Dox. mFR, mean firing rate. b: Regression lines obtained from the relationship between mFR (in spikes/s) per burst and the ETCO2 (%) for the motoneurones illustrated in (a). The slopes of the regression lines represent the neuronal sensitivity or gain to ETCO2 changes (SmFR, in spikes s−1 %−1 ).

8. Bovine serum albumin (BSA) fraction V. 9. Goat-anti-hexon antibody (Biodesign, B65101G). Keep the antibody in aliquots at –20◦ C. 10. Secondary HRP-conjugated antibody: rabbit anti-goat (ZYMED, 81-1620). Keep the antibody at 4◦ C, do not freeze. 11. Diaminobenzidine (DAB) tablet. Keep at 4◦ C, protect from light. 12. Ammonium chloride (NH4 Cl). 13. Ammonium nickel sulphate. 14. Glucose oxidase (Sigma, G-0543). Keep at 4◦ C; protect from light. 2.2. Viral Transduction of NTS or Dorsal Vagal Complex (DVC) in Rats

1. Anaesthesia for viral vector injection into NTS of the rat: Ketamine (60 mg/kg) and medetomidine (250 g/kg) are prepared in 0.9% NaCl and injected intramuscularly (i.m.) (see Note 1). 2. Stereotaxic apparatus: Type: SR-6 N; model no: 98005; Narishige Scientific Instrument Lab, Japan. 3. Syringe pump: Serial No: 207377; model no: Genie; Kent Scientific Corp., USA. 4. Glass capillaries for viral injection. Calibrated microcapillary pipettes (1–5 µL). These pipettes have 1 µL marks which

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make it easy to monitor the volume of the injected virus. The pipettes need to be pulled using a standard pipette puller and their tips broken to ∼20 µm to allow easy flow of the viral suspension. 5. Control of gene expression using a Tet-sensitive viral expression system. For in vitro tests, doxycycline is added to the culture medium as required at the concentration of 2 µg/mL. For in vivo tests, Dox is administered at a concentration of 2 mg/mL supplemented with 5% sucrose into the drinking water (see Note 2). 2.3. Viral Transduction of a XIIth Nerve After a Crush-Induced Injury in Rats

1. Anaesthesia for viral vector injection into XIIth nerve of the rat: chloral hydrate (0.5 g/kg) prepared at 7% in 0.9% NaCl and injected intraperitoneally (i.p.). 2. For virus injection, a Hamilton syringe 7105 N (Part no: 88000) is used. 3. Syringe is connected to the injection glass needle with a vinyl tube (C312VT; Plastics One, USA). 4. Paraffin oil is used to cover the tissues. 5. Glass hooks are used for nerve dissection. 6. Three MM-3 micromanipulators: Narishige Scientific Instrument Lab, Japan. 7. Glass pipettes for viral injection: Borosilicate glass (World Precision Instruments; Item no. 1B200F-6). 8. Injection micropipettes are pulled using a PE-21 puller (Narishige). Their tips are broken to ∼50 µm.

2.4. Lentiviral Vector Production

1. Lenti-XTM 293T cell line (Clontech, 632180). 2. Culture medium and trypsin–EDTA solution for LentiXTM 293T cell line (as described for HEK293 cells in Section 2.1). 3. Lentivirus packaging plasmids pNHP, pCEP4-tat, and pHEF-VSVG and lentivirus transducing plasmids, based on the pTYF backbone into which the gene of interest was cloned (24). 4. SuperFect Transfection Reagent (Qiagen, 301305). 5. 150 mL filter unit with polyethersulfone membrane, 0.45 µm pore size (Fisher, 156-4020). 6. 84 mm Ultracone Centrifuge Tube (Seton Scientific, Part No. 5067). 7. Sucrose analytical reagent grade (Fisher, S/8600/53) prepared as a 20% (w/v) solution in H2 O.

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8. TE671 medulloblastoma cell line. 9. 12-well tissue culture plate. 10. Hexadimethrine bromide at 0.8 mg/mL stock is prepared in sterile PBS and stored at –20◦ C. 11. Paraformaldehyde solution. 12. Levamisol hydrochloride at 50 mM is prepared in water and stored at –20◦ C. 13. 5-Bromo-4-chloro-3-indolyl phosphate (BCIP) at 10 mg/mL prepared in dimethyl formamide and stored at –20◦ C. 14. Nitroblue tetrazolium from Sigma at 50 mg/mL, prepared in water and stored at –20◦ C. 15. BCIP buffer composed of 100 mM Tris base (pH 9.5), 100 mM NaCl, 50 mM MgCl2 made to a total volume of 500 mL. BCIP buffer is stored at 4◦ C. 16. Reaction solution: Combine 6.24 mL BCIP buffer with the addition of 130 µL nitroblue tetrazolium stock, 65 µL levamisol stock and 65 µL BCIP stock. 2.5. Western Blot Analysis

1. Phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2 HPO4 •7H2 O and 1.4 mM KH2 PO4 in ddH2 O, pH 7.4. 2. Radioimmuno precipitation assay buffer (RIPA): 50 mM Tris, 1% nonidet P40, 1% sodium deoxycholate, 0.1% SDS, 150 mM NaCl and 1 mM EDTA in ddH2 O, pH 7.5. Store at 4◦ C. 3. 10% (w/v) nonidet P40. Store at 4◦ C in dark. 4. Homogenizer mortar and pestle. Fisher (1 mL capacity). 5. Protease inhibitor cocktail: 104 mM AEBSF, 0.08 mM aprotinin, 2 mM leupeptin, 4 mM bestatin, 1.5 mM pepstatin A and 1.4 mM E-64 in DMSO. Store in aliquot at –20◦ C. 6. BCA (bicinchoninic acid) protein assay kit: Thermo Scientific Pierce. 7. Laemmli sample buffer (5×): 1.125 M Tris–HCl, 5% (w/v) SDS, 5% (v/v) β-mercaptoethanol, 50% (v/v) glycerol and 0.02% bromophenol blue in ddH2 O, pH 6.8. Add β-mercaptoethanol just prior to use (see Note 3). 8. Separating buffer 1 L: 0.75 M Tris (90.8 g), 0.2% SDS (2.0 g), pH 8.8. Titrate pH and make up to 1 L with ddH2 O. Store at room temperature.

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9. Stacking buffer (1 L): 0.25 M Tris (30.3 g), 0.2% SDS (2.0 g), pH 6.8. Titrate pH and make up to 1 L with ddH2 O. Store at room temperature. 10. 30% Acrylamide/bis solution (see Note 4). 11. APS (ammonium persulphate) (see Note 5). 12. N,N,N′ ,N′ -Tetramethylethylenediamine (TEMED) (see Note 6). 13. 10× Running buffer (1 L): Tris 30 g, glycine 140 g, SDS 10 g, pH 8.3. Titrate pH and make up to 1 L with ddH2 O. Store at room temperature. 14. Prestained protein standard. 15. Bio-Rad Mini Protean II system. 16. PVDF (polyvinylidene difluoride) membrane. 17. 1× Transfer buffer (1 L): Tris 3.45 g, glycine 16.5 g, methanol 100 mL, and make up to 1 L with ddH2 O. 18. 10× Tris-buffered saline (TBS) (500 mL): 0.5 M Tris (30.3 g), 1.5 M NaCl (43.9 g), pH 7.6. Titrate pH and make up to 500 mL with ddH2 O. Store at room temperature. 19. Tween 20 (polyoxyethylene sorbitan monolaurate). Store at ambient temperature. 20. TBS with 0.1% Tween 20 (TBST) buffer: Add 1 mL of Tween 20 to 1 L of TBS, mix and store at room temperature. 21. Polyclonal rabbit anti-nNOS antibody. Store in aliquot at –20◦ C. 22. HRP (horseradish peroxidase)-conjugated anti-rabbit antibody. Store at 4◦ C. 23. Immun-Star Western C chemiluminescent kit containing 50 mL of luminol/enhancer and 50 mL of peroxide solution. Store at room temperature in dark. 24. Autoradiography cassette. 25. Amersham hyperfilm. 26. Film processor: Curix 60, Agfa-Gevaert Group, Germany. 27. Stripping buffer: 62.5 mM Tris, 2% SDS, 100 mM β-mercaptoethanol, pH 6.8. 28. Monoclonal mouse anti-β-actin antibody. Store in aliquots at –20◦ C. 29. HRP (horseradish peroxidase)-conjugated anti-mouse antibody. Store at 4◦ C.

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3. Methods 3.1. Adenoviral Vector Preparation

3.1.1. Amplification

The AVV system is based on human adenovirus serotype 5 subgroup C with the deleted E1 part of the genome. They are amplified in HEK293 cells and purified by double CsCl density gradient centrifugation. The titration protocol described here is an adaptation of the Bewig and Schmidt method (25). It takes advantage of low-level expression of viral hexon proteins in transduced cells. Dilutions of the AVV stock in question are used to infect HEK293 cells. Two days later, these cells are fixed and stained with the antibody specific to the adenovirus hexon protein. Signal is detected after a secondary antibody conjugated with horseradish peroxidase (HRP) amplifies the signal of the antihexon antibody. Subsequent exposure to metal-enhanced DAB substrate turns only the infected cells dark brown. Then the titre of the stock can be determined by counting the number of brown cells in a given area. Each stained cell corresponds to a single ifu. Detailed protocols for AVV amplification and titration are given below.

1. Infect a T75 plate of HEK293 cells (60–80% confluency) with about 2.5 × 108 ifu of virus to be amplified. 2. Harvest the crude viral suspension when complete cytopathic effects (CPE) happens. Store at –20◦ C (see Note 7). 3. Prepare 10 × T150 plates of HEK 293 cells at 80% confluency. 4. Thaw the crude viral suspension harvested from T75 at 37◦ C. Spin the cells down. Keep the supernatant (see Note 8). 5. Infect the 10 × T150 plates with 1 mL viral supernatant obtained from step 4 for each flask. 6. Harvest cells when CPE is complete; this usually takes 2– 3 days. Harvest the cell suspension in Falcon tubes and spin at 200 rcf for 5 min. Keep the cell pellets. Pool the cells together by resuspending in a total volume of 2.5 mL 0.1 M Tris–HCl (pH 8.0) buffer (see Note 9). 7. Freeze and thaw once to break open the cells. 8. Sonicate the viral suspension for 4 min on ice (see Note 10). Pellet cell debris by centrifuging at 200 rcf for 5 min (see Note 11). 9. Transfer cell suspension to UCF tubes, avoiding the cell debris pelleted at the bottom of the Falcon tube. Add 1.8 mL of saturated CsCl (see Note 12) per 3.1

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mL supernatant. Top up tubes with 0.375 vol CsCl (see Note 13). Seal the tubes. Spin in ultracentrifuge 25,000 rcf overnight at 15◦ C (see Note 14). 10. Collect viral band and transfer to a new tube (see Note 15). Top up with 0.375 vol CsCl. Centrifuge in ultracentrifuge at 25,000 rcf for 4–6 h at 15◦ C. 11. Collect viral bands and put through PD-10 columns. Preequilibrate columns with 25 mL of 10 mM Tris pH 7.5, 1 mM MgCl2 . Load 2.5 mL of viral suspension. Add 3.0 mL 10 mM Tris pH 7.5, 1 mM MgCl2 to elute viral protein and collect 2.5 mL of elute (see Note 16). 12. Filter sterilize virus (0.22 µm) and aliquot in small volumes (25 µL). Freeze quickly in liquid nitrogen and store at – 80◦ C (see Note 17). 3.1.2. Titration

1. Plate HEK293 cells (5 × 105 cells/mL) in a 12-well plate in standard culture medium (DMEM + 10% FBS + antibiotics). 2. Using DMEM as the diluent, prepare tenfold serial dilutions of adenoviral stock from 10–2 to 10–8 . 3. Transfect HEK293 cells by adding 100 µL of viral dilution dropwise per well (see Note 18). 4. Incubate for 48 h. 5. Aspirate media and leave the cells to dry in the hood for about 10 min. 6. Fix cells by adding 1 mL/well ice-cold 100% methanol (see Note 19). 7. Place into –20◦ C freezer for 10 min. 8. Aspirate methanol and gently rinse the wells three times with 1 mL PBS + 1% BSA (see Note 20). 9. Take up one 10-µL aliquot of anti-hexon antibody. Dilute in 10 mL PBS + 1% BSA. 10. Aspirate final rinse of PBS + 1% BSA from the wells and add 0.5 mL of hexon antibody dilution to each well. Incubate 1 h at 37◦ C on an orbital shaker. 11. Remove primary antibody, rinse wells 3× PBS + 1% BSA. 12. Dilute secondary HRP-conjugated antibody 1:500 in PBS + 1% BSA and add 0.5 mL to each well. Incubate 1 h at 37◦ C. 13. Meanwhile, prepare 20 mL DAB reaction mix: Dissolve a 10 mg DAB tablet in 10 mL PBS (see Note 21). Then add:

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200 µL 0.4% NH4 Cl (see Note 22). 200 µL 20% glucose. 8.8 mL H2 O. 800 µL 1% ammonium nickel sulphate (see Note 23). Filter through syringe filters into a new foil-wrapped tube. 14. After incubation, remove and rinse the secondary antibody off three times with PBS + 1% BSA. 15. Add DAB reaction mix 0.5 mL/well, wrap plate in foil and incubate at room temperature while gently shaking for about 9 min. 16. Meanwhile, remove 10 mL of the remaining DAB reaction mix and add 20 µL of glucose oxidase. 17. Add 0.5 mL/well of the DAB reaction mix and glucose oxidase for 10 min. Monitor the reaction under a microscope until a dark reaction product appears (see Note 24). Stop reaction by adding excess PBS while background staining is still low. 18. Remove fluid and add fresh PBS. 19. Count stained cells in fields of view at appropriate dilutions (see Notes 25 and 26) and calculate titre in ifu/mL: (average stained cells/field) × (fields/well) volume virus (ml) × (dilution factor) 3.2. Viral Transduction of NTS or DVC in Rats

1. Place anaesthetized animal into a stereotaxic apparatus with the head bent downwards at about 15–20◦ . 2. Make an incision along the midline and the superficial layer of muscle and separate in the middle (see Notes 27 and 28). 3. Pull the neck muscles apart with small tweezers and fit a small wound expander into the wound to keep the muscles apart (see Note 29). 4. Perform injections using a syringe pump and a 50 or 25 µL Hamilton syringe (see Note 30). 5. Administer bilateral microinjections of viral suspension into the NTS at separate sites spanning ± 500 µm rostral/caudal to the calamus scriptorius and 350–700 µm from midline and 500–600 µm below the dorsal surface of the medulla (see Note 31). DVC injection is very similar to NTS injection; for more details please refer to (26).

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3.3. Viral Transduction of a XIIth Nerve After a Crush-Induced Injury in Rats

1. Anaesthetize rats by intraneural injection and surgically expose the right XIIth nerve (see Note 32). 2. Fill 50 µm glass micropipettes with broken tips with a 3 µL solution of the viral suspension. 3. Advance the micropipettes through the perineurium along the axis of the nerve using a micromanipulator towards the branching point. 4. Slowly administer intraneural microinjection of the viral solution for a period of 5 min driven by an oil-filled tubing system connected to a Hamilton syringe. Apply gentle counter-traction to the nerve during the injection. During the same surgical session, crush the nerve at or 10 mm proximal to the bifurcation (27).

3.4. Lentiviral Vector Production

3.4.1. Lentiviral Vector Construction

The LVV system is derived from HIV-1 and pseudotyped with the vesicular stomatitis virus coat. LVV stocks are produced by transient co-transfection of the shuttle plasmids (pTYF backbones in our case), the packaging vector pNHP, Tat plasmid pCEP4-tat and the envelope plasmid pHEF-VSVG in Lenti-X 293T cells. Protocols for viral concentration and titration are modified from Coleman et al. (24). 1. Routinely culture Lenti-X 293T cells in T75 cell culture flasks containing a final volume of 12 mL DMEM full media and incubate at 37◦ C in a humidified atmosphere of 95% air and 5% CO2 . 2. To subculture, aspirate medium, wash cells in 5 mL of DMEM without supplements and incubate with 3 mL trypsin/EDTA until cells dissociate from the culture flask. 3. Mix the solution of dissociated cells with 3 mL DMEM full medium and centrifuge at 120 rcf for 4 min using a standard bench-top centrifuge. Resuspend the cell pellet in 10 mL fresh DMEM full medium (for continued culture, 10% of the resuspended cell pellet is used to inoculate a new T75 flask). For lentiviral production, a confluent T75 flask of Lenti-X 293T cells is required on day 1 and therefore should be inoculated appropriately (see Note 33). 4. Day 1, pm. Dissociate a confluent T75 flask of Lenti-X 293T and resuspend in 26 mL DMEM full medium. Using the cell suspension, inoculate two T150 and one T25 cell culture flask with 12 and 2 mL volumes, respectively. Add DMEM full media to bring final volumes to 20 and 5 mL in the T150 and T25 flasks, respectively (see Note 34). 5. Day 2, am. When Lenti-X 293T are approximately 60% confluent, prepare plasmid and superfect mixes as described in Table 15.1 and vortex for 10 × 1 s pulses.

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Table 15.1 Quantities required for the preparation of the plasmid/ superfect mix Component

2 × T150

1 × T25

pNHP

30 µg

2.5 µg

pHEF-VSVG

12 µg

1 µg

pCEP4-tat

2.5 µg

0.2 µg

pTYF

15 µg

1.25 µg

Serum-free media

1.8 mL

0.1 mL

SuperFect (Qiagen)

120 µL

10 µL

6. Allow the plasmid/superfect mix to stand for 15 min at room temperature. Add 22 mL or 1.8 mL DMEM full medium to the T150 or T25 mixes, respectively. Remove media from the Lenti-X 293T cells and add the plasmid/superfect mixes (12 mL per T150). 7. Incubate the Lenti-X 293T cells at 37◦ C with the plasmid/superfect mix for approximately 6 h. Replace the mix with fresh DMEM full media (12 mL per T150 and 2 mL per T25) and incubate overnight. 8. Day 3, pm. Harvest 1: Collect all medium from virusproducing cells (both 2 × T150 and T25) approximately 24 h following the addition of fresh DMEM full medium, pool together and store at 4◦ C. Add fresh DMEM full medium to the cells (14 mL per T150 and 2.3 mL per T25) and incubate overnight. 9. Day 4, am. Harvest 2: Collect the second harvest medium approximately 18 h after the first harvest medium. Pool both harvests together and filter the medium through a pre-chilled 150 mL 0.45 µm vacuum filter. 10. Using ultracentrifuge tubes add 28 mL filtered viruscontaining medium on top of 0.5 mL 20% sucrose (see Notes 35 and 36). Purify the lentivirus by centrifugation at 10,000 rcf, 4◦ C for 2 h using a Sorvall Discovery 90SE Ultracentrifuge and Sorvall rotor AH-629 (DJB Labcare, Newport Pagnell, UK). 11. Remove all medium and sucrose and add 25 µL sterile phosphate-buffered saline (PBS) to each tube. Tubes are kept at 4◦ C overnight before lentivirus pellets are resuspended and pooled (see Note 37). 12. Day 5, am. Pool 5 µL aliquots of lentivirus and store at –80◦ C (see Note 38).

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3.4.2. Lentiviral Vector Titration

The number of infection particles generated during the lentiviral preparation protocol is determined by staining for PLAP (placental alkaline phosphatase) activity in infected cells. Note that this protocol only estimates the overall quantity of virus production. Titres of LVV with different inserts may vary considerably and this protocol may not reflect it. Other methods such as ELISA or Real-Time PCR are also used routinely and in some cases kits are commercially available. 1. Culture the TE671 cell line as described for the Lenti-X 293T cell line in Section 3.4.1. Points 1–3. 2. Day 1, pm. Dissociate TE671 cells from a confluent T75 flask and purify by centrifugation. Resuspend the cell pellet in 10 mL DMEM full medium and count cells using a haemocytometer (Fisher). Seed a 12-well cell culture plate with 150,000 cells/well in a final volume of 1 mL/well DMEM. 3. Day 2, am. Dilute the hexadimethrine bromide stock 100× in DMEM full medium to a final volume of 10 mL. Remove culture medium from the 12-well plate and add 0.5 mL diluted hexadimethrine bromide to each well. 4. Dilute an aliquot of lentivirus as follows: A: 2 µL virus into 198 µL diluted hexadimethrine bromide (100× dilution). B: 20 µL of A into 180 µL diluted hexadimethrine bromide (1,000× dilution). C: 5 µL of B into 495 µL diluted hexadimethrine bromide (100,000× dilution). 5. In duplicate, add the following volumes of lentivirus dilutions to separate wells: 10 µL A, 10 µL B, 100 µL C, 10 µL C. 6. Day 3, am. Remove the lentivirus-containing media above the TE671 cells 24 h post-infection and replace with 1 mL fresh DMEM full media. 7. Day 4, am. Wash the TE671 cells approximately 48 h postinfection with PBS and then fix the cells by adding 1 mL of 4% paraformaldehyde/PBS to each well. 8. Incubate the cells for 10 min at room temperature and remove the paraformaldehyde by washing three times in PBS. 9. Inhibit endogenous alkaline phosphatase activity by adding 2 mL 75◦ C PBS to each well. Incubate cells at 75◦ C for 1.5 h (PLAP encoded by the lentiviral vector is stable at this temperature).

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10. Dilute levamisol to 500 µM (an endogenous alkaline phosphatase inhibitor) in BCIP buffer. Remove PBS from the TE671 cells and add the levamisol solution (1 mL/well). Incubate at room temperature for 30 min (PLAP is not inhibited by levamisol). 11. Remove levamisol and add 0.5 mL reaction solution to each well. Incubate at room temperature for 1 h and then overnight at 4◦ C (see Note 39). 12. Day 5, am. Infected cells, exhibiting PLAP activity, contain a dark purple precipitate. For each well count the positive cells in six randomly selected fields of view under an inverted light microscope with 10× objective lens (see Note 40). 13. For each well of infected, stained TE671 cells, calculate the viral titre using the following equation (viral titre is determined by taking the average titre derived from individual wells): viral titre (infectious units/mL)=(a∗ b ∗ 1/c)/(d ∗ e) where a = Mean stained cell number per field of view. b = Total fields of view per well (calculated as 157 in the Corning 12-well plate using a 10× objective lens with field of view diameter 1.72 mm). c = PLAP per G.O.I. preparation fraction (in the methodology described in the Lentivirus preparation section, c = 1/12). d = Volume of virus used in infection. e = Dilution factor.

3.4.3. In Vitro LVV Vector Transduction

For testing the activity of the constructs, cell line experiments are required. The in vitro transduction experiments are carried out in PC12 cells. They are grown in DMEM supplemented with 10% heat-inactivated FBS and 5% horse serum. 1. Split PC12 cells and seed in 24-well plates at a cell density of 5 × 104 per well with 0.5 mL culture medium. 2. Transduce cells after 24 h overnight with appropriate LVVs in the presence of polybrene (8 µg/mL). 3. Wash the cells in PBS and culture in full medium for a further 48 h. At the end of incubation, wash the cells and permeabilize with 100 µL of RIPA buffer plus a protease inhibitor cocktail for Western blot analysis as described in Section 3.5. The lysed samples can be kept at –80◦ C until processing.

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3.5. Western Blot Analysis of nNOS Protein

3.5.1. Sample Preparation for Western Blotting Analysis of nNOS

The efficacy of nNOS knock-down can be assessed using Western blot. The nNOS Western blot analysis was carried out as previously described (28, 29). Briefly, total protein was extracted from homogenized samples, followed by quantification with a BCA protein assay kit. Twenty micrograms of total protein per lane were separated on sodium dedecyl sulfate-polyacrylamide gels and transferred to PVDF membranes. The membranes were blocked in 5% non-fat dry milk (NFDM) in TBST for 45 min at room temperature and incubated with polyclonal rabbit antinNOS antibody at 1:5,000 in 3% NFDM-TBST or monoclonal anti-beta-actin antibody at 1:5,000 in 1% BSA-TBST at 4◦ C overnight. Following incubation with polyclonal swine anti-rabbit immunoglobulins/HRP at 1:5,000 in 3% NFDM-TBST or polyclonal rabbit anti-mouse immunoglobulins/HRP at 1:10,000 in 1% BSA-TBST for 90 min, the immunoreactivities were detected with an Immun-Star Western chemiluminescent kit and Amersham high-performance autoradiography film. Bio-Rad Quantity One Software was used to quantitatively compare the relative blots intensities. 1. Harvest fresh rat brain tissues and wash with ice-cold PBS. 2. Homogenize 50 mg of rat brain tissue per sample on ice in 1 mL of RIPA buffer with 5 µL of protease inhibitor cocktail. Use a homogenizer mortar and pestle. 3. Transfer the tissue lysate to 1.5 mL Eppendorf tubes and centrifuge at 14,000×g, 4◦ C for 10 min. 4. Aspirate the total protein extraction supernatant and store at –80◦ C. 5. Quantitate protein concentration using BCA protein assay kit according to the manufacturer’s instructions 6. Mix 20 µg of total protein per sample and 4 µL of Laemmli sample buffer containing β-mercaptoethanol and bring volume to 20 µL with ddH2 O. 7. Boil the resultant mixture in hot-block at 98◦ C for 5 min. 8. Cool the tissue lysate to room temperature for loading.

3.5.2. Electrophoretic Separation (SDS-PAGE)

1. This instruction is based on the use of Bio-Rad Mini Protean II system, but it is easily adaptable to other formats. 2. Wash the large and small glass plates, put spacers right at the edge of the glass plates (see Note 41), and screw the plates and spacers together tightly. When attaching the plate holder to the stand, ensure the side with the heads of the screws is away from you. Make a mark at 1 cm from the top of the small plate.

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3. Prepare a 1.0 mm thick, 10% separating gel by mixing 5 mL of separating buffer with 3.3 mL of 30% acrylamide/bis solution, 1.6 mL ddH2 O, 100 µL of 10% APS and 10 µL TEMED. Immediately load up to 4 mL of separating gel to the level marked on the small plate, leaving the space for the stacking gel (see Note 42). Overlay 500 µL of 75% ethanol to the gel immediately (see Note 43). 4. Let the separating gel set for 20 min. 5. Pour off the 75% ethanol and blot the glass plates dry. 6. Prepare the 5% stacking gel by mixing 5 mL of stacking buffer with 1.6 mL of 30% acrylamide/bis solution, 3.3 mL ddH2 O, 100 µL of 10% APS, 10 µL TEMED. Load the stacking gel from the level marked on the small plate (see Note 44). 7. Place the comb over the space between the two glass plates at an angle and push it into the stacking gel until the two side prongs of the comb are resting on the top of the spacers. 8. Let the stacking gel set for 20 min. 9. Remove the comb carefully and blot the gel top to remove liquid from the wells. 10. Assemble the gel electrophoresis unit. 11. Fill the inner chamber (between the two gels) with running buffer (see Note 45). 12. Pour the rest of running buffer into the outer chamber (see Note 46). 13. Load 5 µL of prestained protein standard to the first well at the edge of the left spacer and 20 µL of samples to be analyzed to the rest wells. 14. Run the gel at 80 V. 15. When the samples pass through the stacking gel, run the gel at 160 V for 1 h. 3.5.3. Western Blotting for nNOS

1. The instructions assume the use of a vertical wet transfer apparatus with PVDF membrane. 2. Pre-wet the PVDF membrane in methanol for 1 min, then submerge the membrane in transfer buffer for 10 min. 3. Disassemble the above gel unit, leave the gel on the small plate and discard the stacking gel. 4. Wet the sponge pads and filter papers in transfer buffer, and make the transfer sandwich by placing on the black piece of the case the sponge pad, filter papers, gel, PVDF membrane, filter papers and sponge pad sequentially. Make

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sure the protein standard is on the right-hand side (see Note 47). 5. Close the transfer sandwich with the white piece of the case, and mount the sandwich in the transfer apparatus, putting the black side of the case near the black side of the transfer apparatus. 6. Remove the cooling block from storage at –20◦ C and place into transfer tank (see Note 48). 7. Fill the transfer tank with transfer buffer and transfer at 250 mA for 2 h. 8. Disassemble the transfer sandwich, remove the PVDF membrane to a shallow dish. 9. Block the membrane with 10 mL of 5% NFDM in TBST buffer at room temperature for 45 min, with gentle shaking at 60 rpm. 10. Wash the PVDF membrane three times in 15 mL of TBST buffer for 10 min, with vigorous shaking at 150 rpm. 11. Incubate the PVDF membrane with polyclonal rabbit antinNOS antibody at dilution of 1:5,000 in 3% NFDM-TBST at 4◦ C overnight, with gentle shaking at 60 rpm. 12. Wash the PVDF membrane three times in 15 mL of TBST buffer for 10 min, with vigorous shaking at 150 rpm. 13. Incubate the PVDF membrane with HRP-conjugated antirabbit antibody at dilution of 1:5,000 in 3% NFDM-TBST for 90 min, with gentle shaking at 60 rpm. 14. Wash the PVDF membrane three times in 15 mL of TBST buffer for 10 min, with vigorous shaking at 150 rpm. 15. Make chemiluminescence HRP detection reagent by mixing 1 mL of peroxide solution and 1 mL of luminol/enhancer provided in the Immun-Star Western C chemiluminescent kit. 16. Incubate the PVDF membrane with the 2 mL of chemiluminescence detection reagent for 5 min at room temperature. 17. Wrap the PVDF membrane with cling film, place into an autoradiography cassette and develop in a dark room. 18. Expose the PVDF membrane to Amersham hyperfilm for 1 min (see Note 49) and develop in a film processor to display the nNOS blot at 160 kDa. 19. After chemiluminescence development, wash the PVDF membrane three times in 15 mL of TBST buffer for 10 min, with vigorous shaking at 150 rpm.

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20. Incubate the PVDF membrane in stripping buffer in a heat-sealed plastic bag at 50◦ C for 30 min with gentle agitation. 21. Wash the PVDF membrane three times in 15 mL of TBST buffer for 10 min, with vigorous shaking at 150 rpm. 22. Block the membrane with 10 mL of 5% NFDM in TBST buffer at room temperature for 45 min, with gentle shaking at 60 rpm. 23. Wash the PVDF membrane three times in 15 mL of TBST buffer for 10 min, with vigorous shaking at 150 rpm. 24. Incubate the PVDF membrane with monoclonal mouse anti-β-actin antibody at dilution of 1:5,000 in 1% BSA-TBST at 4◦ C overnight, with gentle shaking at 60 rpm. 25. Wash the PVDF membrane three times in 15 mL of TBST buffer for 10 min, with vigorous shaking at 150 rpm. 26. Incubate the PVDF membrane with HRP-conjugated antimouse antibody at dilution of 1:10,000 in 1% BSA-TBST for 90 min, with gentle shaking at 60 rpm. 27. Wash the PVDF membrane three times in 15 mL of TBST buffer for 10 min, with vigorous shaking at 150 rpm. 28. Repeat chemiluminescence development as described for nNOS from Steps 15–18 to display β-actin blot at 42 kDa. 29. Scan the film and quantify the blots of nNOS and β-actin by densitometry using Quantity One software.

4. Notes 1. Always freshly prepare anaesthetics. 2. Supply fresh Dox every 3 days. 3. The β-mercaptoethanol and SDS powder are hazardous. Prepare solution in a ventilated fume hood. 4. The acrylamide/bis solution is neurotoxic when unpolymerized, so care should be taken not to receive exposure. 5. APS may cause fire and irritation to eyes, respiratory system and skin. 6. TEMED is highly flammable, and is harmful if inhaled or ingested. 7. Crude viral suspension includes cells and growth media. When CPE happens, cells round and lift off from the culture flasks.

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8. Most viral particles are now released to supernatant. 9. The majority of viral particles are kept in the cells. 10. Between each minute of sonication, mix the sample well and leave for about 2 min to make sure it is ice cold before the next round of sonication. 11. Handle the tubes carefully as the cell pellet can get easily dislodged. 12. Be sure the saturated CsCl stock is equilibrated to room temperature prior to use as temperature affects concentration. 13. 0.375 vol CsCl is prepared by mixing 6 mL saturated CsCl with 10 mL 0.1 M Tris–HCl, pH 8.0. 14. Vortex thoroughly before centrifugation. 15. Collect viral band by puncturing the top of the tube with a 25 Gauge needle and puncturing the bottom with another needle attached to a 5 mL syringe. Take out the viral band by pulling the syringe. 16. Discard the first 0.5 mL as this fraction does not contain virus. 17. Avoid repeated freeze–thaw cycles as this causes a dramatic decrease of viral titres. 18. The degree of error introduced in each serial dilution may affect the result, so perform duplicate infections to ensure accurate assay results. 19. Add the fixative very gently taking care not to dislodge cell monolayer. 20. Avoid dislodging the cell monolayer. 21. Use a magnetic stirrer in an aluminium-foil-wrapped 50 mL tube for 5–10 min. The tablet may not dissolve totally. 22. Solution will last in fridge for 2–3 weeks. 23. Solution will last in fridge for 2–3 weeks. 24. Colour development should take 1–2 min. 25. Count 2–3 adjacent cells as one. 26. It is important that the counted fields be selected in an unbiased manner. We recommended that a minimum of three fields are selected to count and that the counted fields contain 10–50 positive cells – assuming that the distribution of infected cells is random over the entire well. 27. Make a longitudinal midline incision of the skin beginning from lower part of the skull for 2 cm. Only cut the most superficial layer of muscle to prevent bleeding. Muscles underneath are separated along the midline with forceps

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and a small wound expander can be fitted to keep them separate. 28. To open the membrane between the skull and the first vertebrum the edge of a fresh fine needle can be used as a miniature blade. 29. Perform Notes 29, 30 and 31 under a long-range binocular microscope. 30. A syringe is connected to the pipette using fine plastic tubing. Use a piece of soft silicone tubing to connect the glass pipette. The tubing can be softened by placing into petroleum ether for 15–30 min. 31. To avoid post-surgical cardiovascular effects, animals should be allowed 7 days to recover before blood pressure data is sampled. 32. After the lesion procedure the nerve became translucent, confirming that transection did not occur. 33. Lenti-X 293 cells should be seeded appropriately to achieve around 100% confluence at day 1 of the lentiviral production protocol, and seeding density may be calculated assuming a cell doubling time of 24 h under the described culturing conditions. 34. The two T150 flasks are used for the generation of lentivirus containing the gene of interest (G.O.I.), whereas the T25 is used for the production of lentivirus containing placental alkaline phosphatase (PLAP) driven by the human elongation factor-1α promoter. The latter is required for titration of the lentivirus batch. 35. Care should be taken when overlaying virus-containing medium above sucrose. To achieve a clean interface between medium and sucrose layers, add 10 mL virus containing medium to the centrifuge tube and subsequently pipette the sucrose directly into the bottom of the tube. Proceed to carefully overlay the remaining medium, adding equal volumes to pairs of tubes. 36. Ultracentrifuge buckets containing filled tubes should be weighed before centrifugation. A weight difference of