Nitric Oxide: Methods and Protocols

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

v

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

ix

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

1

1.

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

S ECTION 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

17

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

27

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

39

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

57

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

73

7.

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

9.

81

91

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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Contents

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 S ECTION 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-R OTHKAMM • 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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x

Contributors

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. T IMOSHIN • 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

xi

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

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.

Nitric Oxide Physiology and Pathology

3

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

Nitric Oxide Physiology and Pathology

5

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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Section I Detection and Quantification of Nitric Oxide

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

21

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 .

22

Rothkamm and Burdak-Rothkamm

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.

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

23

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.

24

Rothkamm and Burdak-Rothkamm

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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5.

6.

7.

8.

9.

10.

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.

13.

14.

15.

16.

17.

18.

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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

27

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Nagababu and Rifkind

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 <0.1 μM. The ozone-based chemiluminescence method using Sievers NOA 280i instrument is one of the highly sensitive methods to determine nanomolar concentrations of NO. The details of the instrument and techniques for measurement of NO have been reviewed (8–10). The bond between sulfur and NO must be cleaved in order to determine RSNOs by this method. Several methods are available in the literature for this purpose. They include photolysis (11), Cu(I)-Iodide (12), CuCl2 + cysteine (13), tri-iodide (7), and Cu(II)+ ascorbate (10, 14) to cleave the RSNO bond at a neutral pH or an acidic pH. The KI/I2 reductive reagent has been widely used to determine nitrite and RSNOs in biological fluids (10, 15). It has been shown that this reagent measures the nitrite and N-nitrosamines in addition to RSNOs (16). Marley et al. (12) have employed pretreatment of the sample with acidified sulfanilamide to remove the nitrite interference (12). We developed a method to determine the RSNO using Cu+2 -ascorbic acid at a neutral pH without interference of nitrite (14). This method has a high sensitivity to measure both high and low molecular weight RSNOs including S-nitrosohemoglobin and albuminSNO. However, this method is not suitable for high protein content samples due to formation of foam at neutral pH in the reaction vessel. Since the RSNOs levels in biological samples are in low nanomolar concentrations, a large volume, which contains a high level of protein, needs to be injected to see a signal. Therefore, this method has been modified by using Cu(II)Cl2 and ascorbate in strong acidic acid instead of neutral pH. We found that Cu(II) in the presence of ascorbate (ASC) readily cleaves the RSNO bond to stoichiometrically releases NO. This

Determination of S-Nitrosothiols by Chemiluminescence

29

reagent does not release NO from nitrated lipids, proteins, and N-nitrosomines. Cu(II) + ASC→Cu(I) + ASC• RSNO + Cu(I)→RS− + NO + Cu(II) The released NO in the reaction vessel is carried by inert gas to the detector where it reacts with ozone to produce a chemiluminescence signal proportional to the concentration. This method also detects nitrite in addition to RSNOs. This nitrite interference is eliminated by pretreating the sample with acidified sulfanilamide, which forms a diazonium complex with nitrite that does not generate a chemiluminescence signal.

2. Materials 2.1. Specific Equipment

1. Sievers Nitric Oxide Analyzer model 280 (GE Analytical Instruments, Boulder, CO) 2. Liquid program (3.21) 3. Origin program 4. Oxygen and inert gases (argon or nitrogen) 5. Hamilton syringes 50, 100, 250 μL, with 5 in. long needles 6. Portable centrifuge

2.2. Reagents

1. Water: Ultra pure water (UPH2 O) (at 18.2 M-cm) from PURELAB Plus or Millipore H2 O Milli Q-Gard 2 Purification pack or doubled distilled water (See Note 1). 2. 50 mM diethylenetriaminepentaacetic acid (DTPA) solution: Suspended 983.4 mg DTPA in 25 mL of UPH2 O. Add 1 M NaOH drop wise until it dissolves. Adjust the pH to 7.0. Make up the final volume to 50 mL with UPH2 O. This solution is stable at room temperature. 3. 5% Acidified sulfanilamide: Dissolve 5 g sulfanilamide in 100 mL 2 M HCl. 4. Preparation of S-nitrosoglutathione (GSNO) standard solution. A. Stock solution: 10 mM glutathione solution is prepared by dissolving 30.732 mg of glutathione in 10 mL of 1 M HCl. Similarly, 10 mM nitrite solution is prepared by dissolving 6.9 mg sodium nitrite in 10 mL of UPH2 O containing 0.2 mM DTPA solution (40 μL of 50 mM DTPA diluted to 10 mL with UPH2 O). 5 mL of glu-

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tathione solution is mixed with 5 mL of nitrite solution. This solution should be capped, protected from light, and kept cold (4◦ C). The concentration of this solution is determined by diluting 0.25 mL of GSNO stock solution to 2.5 mL with UPH2 O in a cuvette and an absorption spectrum is run from 250 to 450 nm. The maximum absorbance at 335 nm is recorded. The concentration of GSNO is determined by using the millimolar extinction coefficient of 0.980 at 335 nm. Calculation = Absorbance at 335 nm/0.980 × 10 (dilution factor). This value is in the range of 4–5 mM. The concentration of GSNO is adjusted to 1 mM by appropriate dilution with 0.5 M HCl. B. Working Standard Solution (1 μM): 10 μL of stock solution (1 mM) is diluted in 10 mL of sulfanilamide solution (9:1 V/V, UPH2 O: 5% acidified sulfanilamide). This 1 μM working standard is serially diluted with acidified sulfanilamide (9:1 V/V, UPH2 O: 5% acidified sulfanilamide) at a ratio of 1:2, 1:4 down to a concentration of 7.8 nM (Table 3.1). (See Notes 2, 3, and 4)

Table 3.1 Preparation of GSNO standards

Ratio

Volume of 1 μM GRSNO std (mL)

Volume of acidified sulfanilamide (mL)

Total volume (mL)

Final nitrite concentration (nM)

1:0

10

0

10

1,000

1:2

5

5

10

500

1:4

2.5

7.5

10

250

1:8

1.25

8.75

10

125

1:16

0.625

9.375

10

62.5

1:32

0.3125

9.6875

10

31.25

1:64

0.156

9.844

10

15.625

1:128

0.078

9.922

10

7.812

5. 250 mM N-ethylmaleimide (NEM) solution: Dissolve 313 mg in 10 mL PBS. This reagent should be prepared fresh every day (see Note 5) 6. 100 mM potassium ferricyanide: Dissolve 329 g in 10 mL UPH2 O. 7. RSNOs preservation solution: 9.75 mL potassium ferricyanide mixed with 0.1 mL of triton X-100, 0.1 mL of 250 mM NEM, and 50 μL of 50 mM DTPA.

Determination of S-Nitrosothiols by Chemiluminescence

31

8. 100 mM cupric chloride (CuCl2 ): Dissolve 170.48 mg cupric chloride dihydrate in 10 mL UPH2 O. 9. 125 mM ascorbic acid solution: Dissolve 220.15 mg in 10 mL of UPH2 O. Prepare fresh reagent every day.

3. Methods 3.1. Calibration Curve for RSNOs

1. Add 7.5 mL glacial acetic acid, 200 μL of 100 mM CuCl2, and 200 μL of 125 mM ascorbic acid into the purge vessel. 2. Gas bubbler/sodium hydroxide trap is connected between the purge vessel and the detector. 3. Water bath temperature for circulating water around waterjacked purge vessel is set at 37◦ C. Cold water circulation for condenser is not required. 4. Liquid program is used to acquire data. The reaction mixture is continuously purged with inert gas at a constant cell pressure by adjusting the gas flow. The data is acquired every 0.25 s. Once the baseline is stabilized, 100 μL of working standard solutions from 7.812, 15.62, 31.25, 62.5, 125, 250, 500 and 1,000 nM GSNO is injected into the purge vessel using a 5 in. long needle (see Note 4). Chemiluminescence signals are recorded. Duplicate measurements are made for each sample. 5. Origin program is used to analyze the data: It can be purchased from OriginLab Corporation, One Roundhouse Plaza, Suite 303, Northampton, MA 01060 USA. 6. The data from the liquid program is imported into the Origin program in an ASCII format. Chemiluminescence signals were regenerated by plotting the time (X axis with each row equivalent to 0.00417 min) against mV values (Y axis) (Fig. 3.1a). 7. The area under the curve of each signal is calculated after determination of the baseline and finding the peaks using the tool menu of the program (Table 3.2). 8. Standard Curve: The standard curve is constructed from the average area of the peaks versus concentration. The slope and intercept of the standard curve are determined (Fig. 3.1b).

3.2. Blood Collection

Venous blood is drawn from fasting human subjects into 5 mL heparin vacutainers. 6.5 mM NEM (26 μL: 250 mM NEM/mL blood) and 0.2 mM DTPA (5 μL: 50 mM DTPA/mL blood) are

Nagababu and Rifkind

B

70

18

60

15 Slope= 0.01601 Area (A.U.)

A

50 40 30 20 5

25 0 50 0 10 0 nM 0

12

0 −5 0

12

R=0.99936 Intercept= − 0.025

9 6 3

10 7. 8 15 .6 31 .2 62 .5

Chemiluminescence (mV)

32

5 10 15 20 25 30 35

0 0

200 400 600 800 1000

Time (min)

GSNO (nM)

Fig. 3.1. A. Chemiluminescence signals of GSNO standards: GSNO (7.812–1,000 nM; 100 μL) is injected into the purge vessel of the nitric oxide analyzer which contains 7.5 mL glacial acetic acid, 200 μL of 100 mM CuCl2 , and 200 μL of ascorbic acid. B. Calibration curve for RSNOs is obtained by integrating the area under the curve.

Table 3.2 Average area under the curve of each signal. The area of the blank value (water) 0.201 is subtracted from the area of all the nitrite standards Nitrite concentration (nM) 7.812

Signal area under curve (AU) 0.150

15.625

0.32

31.25

0.467

62.5

0.975

125

1.835

250

4.035

500 1,000

7.89 16.1

AU: Arbitrary Units.

immediately mixed with whole blood and after 1 min, centrifuged at 4,500 rcf for 5 min. NEM and DTPA prevent the decay of plasma RSNO to nitrite (see Notes 2 and 5). Process the plasma and red blood cells as soon as possible after separation from the blood. A. Processing of Plasma: 0.9 mL plasma that should be free of hemolysis is immediately transferred into light protected (dark colored) microtubes that contained 0.1 mL of 5% acidified sulfanilamide (see Note 6). Plasma is immediately frozen on dry ice and

Determination of S-Nitrosothiols by Chemiluminescence

33

stored at –85◦ C or –150◦ C for future analysis or measured immediately. B. Processing of RBCs: 1. 1 mL of RBCs (pipette from the bottom of tube) is mixed with 1 mL of preservation solution and incubated for 5 min to lyse the cells and oxidize the hemoglobin (see Note 7). 2. 0.25 mL of hemolysate is passed through a G-25 Sephadex column, which is pre-equilibrated with 0.1 mM DTPA-PBS buffer, and the hemoglobin fraction is collected. 3. This column separation is carried out under subdued light preferably in a cold room (at 4◦ C). 4. The concentration of methemoglobin is determined using the millimolar extinction coefficient of 3.7 at 630 nm. Concentration of heme (mM) = Absorbance at 630 nm/3.7×dilution factor. The concentration of the heme will be around 2.0 mM. 5. 0.9 mL of this solution is mixed with 0.1 mL of 5% acidified sulfanilamide in 1 M HCl. This sample can be frozen at –85◦ C or –150◦ C for later analysis or measured immediately. 3.3. Measurement of Plasma RSNOs

1. Thaw the plasma under subdued light just prior to determination. Thawed plasma samples are kept on ice until injected (see Note 8). 2. 1 mL plasma sample is divided into two aliquots. One aliquot is incubated with 5 mM HgCl2 (25 μL: 100 mM HgCl2 ) for 10 min and to the second aliquot 25 μL UPH2 O is added. 3. 100 μL plasma is injected using a Hamilton syringe into the bottom of the purge vessel that contains 7.5 mL glacial acetic acid, 200 μL of 100 mM CuCl2 , and 200 μL of 125 mM ascorbic acid (see Note 9). 4. Each sample is injected in triplicate. The reagents are changed after every three or four injections. 5. Data is transferred to the Origin program and the chemiluminescence signals are regenerated as mentioned for the nitrite standard curve. 6. The signals are smoothed by averaging the data of 20 points to eliminate the noise and for baseline correction (Fig. 3.2). 7. Open the smoothed data worksheet and the chemiluminescence signals are regenerated. The area under the curve is determined as described for the standard curve. An

Nagababu and Rifkind

A

B

10

9.0

9

Signals (mV)

Signals (mV)

8

0.74167 2.44583

8.5

+ HgCl2 5.83333 4.17917

8.0 7.5

7 −1 0 1 2 3 4 5 6 7 Time (min)

7.0

−1 0 1 2 3 4 5 6 7 Time (min)

Fig. 3.2. Chemiluminescence signal of plasma RSNOs: 100 μL of plasma treated with 6.5 mM NEM, 0.2 mM DTPA, and 5% sulfanilamide in 2 N HCl as mentioned in methods section is injected into the purge vessel. A. Chemiluminescence signals from the raw data. B. Chemiluminscence signals from the smoothed data using origin program for calculating the area under the curve.

100 S-nitrosothiols (nM)

34

80 60 40 20 0 Plasma

RBCs

Fig. 3.3. RSNOs values in plasma and red blood cells.

example of raw data before and after being smoothed is shown in Fig. 3.2. 8. The RSNOs values are calculated by taking the difference in the area of the chemiluminescence signals for samples in the presence and absence of HgCl2 (see Fig. 3.3). Calculation: An example for a sample of plasma RSNOs (nM): 0.085 – (–0.025)

0.1 mL

0.01601

0.1 mL

1.030 mL

1.0 mL

1.0 mL

0.9 mL

0.95 mL

—————— × ——— × ——— × ——— × ——— = 6.4 nM 1 mL

Numerators = 0.085 = sample area, 0.025 = intercept, 0.1 mL = Std. injected, 1.030 mL = blood + BNEM + DTPA, 1.0 mL = plasma + sulfanilamide, 1.0 mL = plasma + sulfanilamide + HgCl2 Denominators = 0.01601 = slope, 0.1 mL = plasma injected, 1.0 mL = blood volume, 0.9 mL = plasma volume, 0.95 mL plasma + sulfanilamide

Determination of S-Nitrosothiols by Chemiluminescence

3.4. Measurement of RBCs RSNOs

35

1. Thaw the RBC lysate under subdued light just prior to determination. Thawed lysates are kept on ice until injected. 2. 1 mL of freshly processed RBC sample or thawed sample is divided into two aliquots. 5 mM HgCl2 (25 μL: 100 mM) is added to one aliquot and 25 μL UPH2 O is added to the second aliquot and incubated for 5 min. 3. 100–200 μL sample is injected using a Hamilton syringe into the bottom of the purge vessel that contains 7.5 mL of glacial acetic acid, 200 μL of 100 mM CuCl2 , and 200 μL of 125 mM ascorbic acid. 4. Reagents are changed every two injections. 5. The signals are processed as mentioned in Section 3.3 for plasma samples. 6. The RSNO values are calculated by taking the difference in the area of the chemiluminescence signals for samples in the presence and absence of HgCl2 (Fig. 3.3). Calculation: An example for a sample of RBC high molecular weight RSNOs (nM): 0.206 –(–0.025)

0.1 mL

0.01601

0.2 mL

1.0 mL

1.0 mL

20

0.95 mL

1.85

—————— × ——— × ——— × ——— × ——— = 91.2 nM 0.9 mL

Numerators: 0.206 = area, 0.025 = intercept, 0.1 mL = std. injected, 1.0 mL = hemolysate + sulfanilamide, 1.0 mL = hemolysate + sulfanilamide + HgCl2, 20 = heme concentration of RBCs Denominators: 0.01601 = slope, 0.2 mL = hemolysate injected, 0.9 mL = hemolysate, 0.95 mL = hemolysate + sulfanilamide, 1.85 = heme concentration of hemolysate 3.5. Determination of RSNOs in Cells

After completion of the incubation period for cells, the formation of RSNOs is stopped by adding five- to ten fold excess of NEM and allowing 5–10 min to block all the thiol groups (see Note 10). Cells are lysed by sonication and the protein content was determined. Treat the cell lysate with acidified sulfanilamide (9:1 V/V, cell lysate: 5% acidified sulfanilamide) to remove the nitrite and to stabilize the RSNOs. Inject 1–5 mg protein samples into the purge vessel dependent on the RSNOs content (see Note 11).

4. Notes 1. Always use fresh high-quality water (UPH2 O with resistivity = 18.2 M-cm) containing a metal chelator preferably 0.1 mM DTPA to prepare the solutions.

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2. RSNOs are light and temperature sensitive. Samples should be protected from light by carrying out all steps under subdued light and/or using dark color tubes. Keep samples on ice or in a cold room. 3. Make sure that there is no gas leak in the purge system. Maintain constant instrument cell pressure by adjusting the inert gas flow. Run the system at least 30 min for stabilization prior to initiating sample analysis. 4. We strongly suggest the use of the Origin program for integration of peaks. It can be purchased from Origin Lab Corporation, One Roundhouse Plaza, Suite 303, Northampton, MA 01060 USA. 5. RSNOs are unstable in presence of free thiol groups and metal contaminants. In order to preserve the RSNOs, blood should be collected in NEM and DTPA containing vacutainer tubes or NEM and DTPA should be added to blood as soon as it is collected. Allow 1–2 min to react NEM with thiols and centrifuge the blood immediately. 6. RSNOs are more stable under acidic conditions. Transfer the plasma to tubes containing acidified sulfanilamide solution (9:1 V/V, plasma: 5% acidified sulfanilamide) immediately after centrifugation. This reagent removes the nitrite and stabilizes the RSNOs under frozen storage conditions. 7. Thiol groups also should be blocked with NEM before addition of potassium ferricyanide (preservation solution) to RBCs to oxidize the hemoglobin. Otherwise oxidation of iron nitrosylhemoglobin by potassium ferricyanide generates S-nitrosohemoglobin. Addition of NEM together with potassium ferricyanide does not completely prevent the generation of S-nitrosohemoglobin. 8. Thaw the plasma samples just prior to use, while protecting from light. GSNO standards also should be prepared just prior to use. Complete the analysis within 2–3 h. 9. Inject the samples at the bottom of the purge vessel using a syringe with a 5-in. long needle for good reproducibility. Syringe needle should be rinsed with water and wiped after every injection. Otherwise sample will be contaminated with Cu(I) reagent, which degrades RSNOs. 10. Make sure that free thiols are blocked with NEM before the addition of acidified sulfanilamide to the plasma to remove the nitrite. Otherwise free thiols react with nitrite under acidic conditions generating RSNOs. Some investigators have used NEM together with acidified sulfanilamide to block the thiols and remove the nitrite. However,

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we observed that nitrite reacts with thiols to form RSNOs under acidic conditions before it complexes with sulfanilamide. 11. When injecting high protein samples (plasma) leave a sufficient gap between injections until the gas bubbles are regenerated in the purge vessel. References 1. Hogg, N. (2002) The biochemistry and physiology of S-nitrosothiols. Annu Rev Pharmacol Toxicol 42, 585–600. 2. Kharitonov, V. G., Sundquist, A. R., Sharma, V. S. (1995) Kinetics of nitrosation of thiols by nitric oxide in the presence of oxygen. J Biol Chem 270, 28158–28164. 3. Ignarro, L. J., Edwards, J. C., Gruetter, D. Y., Barry, B. K., Gruetter, C. A. (1980) Possible involvement of S-nitrosothiols in the activation of guanylate cyclase by nitroso compounds. FEBS Lett 110, 275–278. 4. Foster, M. W., McMahon, T. J., Stamler, J. S. (2003) S-nitrosylation in health and disease. Trends Mol Med 9, 160–168. 5. Gaston, B. M., Carver, J., Doctor, A., Palmer, L. A. (2003) S-nitrosylation signaling in cell biology. Mol Interv 3, 253–263. 6. Foster, M. W., McMahon, T. J., Stamler, J. S. (2003) S-nitrosylation in health and disease. Trends Mol Med 9, 160–168. 7. MacArthur, P. H., Shiva, S., Gladwin, M. T. (2007) Measurement of circulating nitrite and S-nitrosothiols by reductive chemiluminescence. J Chromatogr B Analyt Technol Biomed Life Sci 851, 93–105. 8. Dunham, A. J., Barkley, R. M., Sievers, R. E. (1995) Aqueous nitrite ion determination by selective reduction and gas phase nitric oxide chemiluminescence. Anal Chem 67, 220–224. 9. Nagababu, E., Rifkind, J. M. (2007) Measurement of plasma nitrite by chemiluminescence without interference of S-, N-nitroso and nitrated species. Free Radic Biol Med 42, 1146–1154.

10. Basu, S., Wang, X., Gladwin, M. T., KimShapiro, D. B. (2008) Chemiluminescent detection of S-nitrosated proteins: comparison of tri-iodide, copper/CO/cysteine, and modified copper/cysteine methods. Methods Enzymol 440, 137–156. 11. Stamler, J. S., Jaraki, O., Osborne, J., Simon, D. I., Keaney, J., Vita, J., Singel, D., Valeri, C. R., Loscalzo, J. (1992) Nitric oxide circulates in mammalian plasma primarily as an S-nitroso adduct of serum albumin. Proc Natl Acad Sci USA 89, 7674–7677. 12. Marley, R., Feelisch, M., Holt, S., Moore, K. (2000) A chemiluminescense-based assay for S-nitrosoalbumin and other plasma S-nitrosothiols. Free Radic Res 32, 1–9. 13. Fang, K., Ragsdale, N. V., Carey, R. M., MacDonald, T., Gaston, B. (1998) Reductive assays for S-nitrosothiols: implications for measurements in biological systems. Biochem Biophys Res Commun 252, 535–540. 14. Nagababu, E., Ramasamy, S., Rifkind, J. M. (2006) S-nitrosohemoglobin: a mechanism for its formation in conjunction with nitrite reduction by deoxyhemoglobin. Nitric Oxide 15, 20–29. 15. Yang, B. K., Vivas, E. X., Reiter, C. D., Gladwin, M. T. (2003) Methodologies for the sensitive and specific measurement of S-nitrosothiols, iron-nitrosyls, and nitrite in biological samples. Free Radic Res 37, 1–10. 16. Rassaf, T., Bryan, N. S., Kelm, M., Feelisch, M. (2002) Concomitant presence of N-nitroso and S-nitroso proteins in human plasma. Free Radic Biol Med 33, 1590–1596.

Chapter 4 Measurement of Nitrite in Blood Samples Using the Ferricyanide-Based Hemoglobin Oxidation Assay Barbora Piknova and Alan N. Schechter Abstract Nitrite is currently recognized as a biomarker of the state of nitric oxide metabolism. Therefore, assessing nitrite levels in various organs and compartments is an important issue. As nitrite levels in most organs and tissues are low (in high nanomolar or low micromolar range) several new sensitive methods for quantifying nitrite in various biological samples have been developed. Chemiluminescence, combined with triiodide reducing solution, is currently considered the most sensitive method, allowing quantification in the low nanomolar range of nitrite concentrations. Here, we present an overview of chemiluminescencebased determination of nitrite in blood and blood compartments – red blood cells and plasma. We also explain how to preserve the original physiological nitrite concentration in nitrite-hostile environments, such as an excess of hemoglobin in blood. Key words: Nitrite, hemoglobin, red blood cell, chemiluminescence.

1. Introduction Recently, nitrite has emerged as a central component of the nitric oxide (NO) cycle – it is both a NO precursor and a product of NO oxidation. This dual role not only makes it a prime target for therapies to correct NO deficiency but also allows insights into NO metabolism and its changes in health and disease. It is thought that simple measurement of nitrite levels over time under different conditions and in different organs/biocompartments can give a rough estimation of NO pathway activity and efficiency. Attempts have been made to use nitrite as a biomarker of some health conditions. The most common clinical measurements of nitrite are those in blood and urine, due to easy access to both fluids during clinical H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_4, © Springer Science+Business Media, LLC 2011

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examination of a patient. Here we will describe measurement of nitrite in blood and its compartments (red blood cells (RBC) and plasma) using the gas phase chemiluminescence method with triiodide reducing solution and a nitrite-preserving “stop” solution, based on original published protocols (1–3). Major steps in necessary pretreatment of blood and sample processing are outlined in Fig. 4.1. Final nitrite concentration determination procedures are outlined in Fig. 4.2.

Fig. 4.1. Outline guide to blood processing for nitrite determination in whole blood, red blood cells (RBC), and plasma.

2. Materials 2.1. NitritePreserving “Stop” Solution

Nitrite rapidly reacts with oxy- and deoxy hemoglobin (Hb) in blood and is destroyed in both reactions (see Note 1). Standard pretreatment of blood and most biological samples, to prevent nitrite destruction, consists of oxidizing the heme to form metHb

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Fig. 4.2. Outline guide to sample measurements and data processing for nitrite determination by chemiluminescence method (CL). Samples prepared as shown in Fig. 4.1.

or heme(FeIII )-protein. This pretreatment is not necessary if the sample does not contain Hb or other heme(FeII )-proteins and it is usually skipped for plasma nitrite measurement as shown in Fig. 4.1. The roles of the individual reagents in the stop solution are explained in Note 2. 2.1.1. Reagents

K3 Fe(CN)6 (potassium ferricyanide, MW 329.24), NEM (N-ethylmaleimide, MW 125.12) – light sensitive, store at 4◦ C, Nonidet P-40 (octyl phenoxylpolyethoxylethanol), H2 O – we recommend using either molecular biology grade water or fresh distilled water filtered using Milli-Q (Millipore, Billerica, MA) water purification system with conductivity of 18 M cm (see Note 3).

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2.1.2. Preparation

Prepare the solution containing 890.9 mM potassium ferricyanide and 118.13 mM NEM. This solution needs to be vortexed vigorously for several minutes until all NEM and ferricyanide crystals are dissolved and the color is clear yellow. Add NP-40 in a 1:9 ratio (v/v, NP-40/solution), and gently mix ingredients together by inverting the test tube several times. At this stage, avoid unnecessary mixing, as this will cause excessive foaming (see Notes 4 and 5).

2.2. Sample Collection and Pretreatment

Blood is collected using a 20 G needle for venous blood and an 18 G needle for arterial blood to prevent hemolysis. 5 IU heparin/mL blood is used to prevent coagulation. Note 6 emphasizes the crucial points in sample collection and processing, and Fig. 4.1 provides a general overview of the procedures involved.

2.3. Chemiluminescence 2.3.1. Chemiluminescence Principle, Reagents, and Reactions

Only free NO gas is detected by chemiluminescence. In order to detect NO metabolites, they have to be converted into free NO prior to quantification. Several reducing/oxidizing solutions, depending on the nature of the NO metabolite, are used for this purpose (see the following paragraph for details about nitritereducing solutions). For a simplified presentation of the chemiluminescence setup see Fig. 4.3 or the more detailed descriptions in (4, 5). Briefly, free NO gas is purged from the reaction vessel by an inert carrier gas (He, N2 , Ar) into the chemiluminescence analyzer. In the analyzer reaction chamber, ozone (O3 ) is combined with NO to form nitrogen dioxide (NO2 ) in its activated state (see reaction 5 in Note 7). Upon deactivation, NO2 ∗ emits a photon (see reaction 6 in Note 7) in the infrared region that is detected by photomultiplier equipped with a long pass filter, detecting only emission above 600 nm. The intensity of emitted light is proportional to the concentration of NO in the reaction chamber, and using proper calibration curves, it can be related to the amount of NO in the original sample (see Note 7). All NO metabolites and adducts, such as nitrite, nitrate, R-nitrosothiols (R-SNO), R-nitrosoamines (R-NNO), or metalNO, must be converted into free NO gas in order to quantify their original amounts via chemiluminescence. A reducing solution is used for nitrite, nitrate, R-SNO, and R-NNO. Here we will focus on the most popular – the I3 -based reducing solution (see Note 8). I3 -based reducing mixture is not specific for nitrite, and all NOx metabolites, except nitrate, will be reduced to NO, contributing to the signal. Details about how to account for this “false” nitrite signal using acid sulfanilamide (AS) treatment are in Note 9.

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Fig. 4.3. Chemiluminescence apparatus setup. Reaction vessel is filed with I3 solution with He carrier gas gently bubbling though. Sample is injected using Hamilton syringe through septum into I3 solution where NOx components are reduced to NO gas and carried into NO analyzer. Cold trap, NaOH-filled trap, and filter protect analyzer against humidity and acid vapors. In reaction chamber (RC) NO gas is combined with O3 (generated in O3 generator) from O2 either from O2 tank or from room air. Chemiluminescence signal from NO2 ∗ is detected by photomultiplier tube (PMT) and further amplified and processed. Data acquisition and analysis are carried out on PC. Vacuum pump created low pressure in the reaction chamber (RC) and evacuates toxic NO2 gas after chemiluminescence measurement through charcoal filter (CF).

1. Reagents for I3 solution: KI (potassium iodide, MW 166.0) – light sensitive, I2 (crystalline iodide, MW 253.81) – light sensitive, glacial acetic acid, H2 O. 2. Prepare 301.2 mM KI together with 137.8 mM I2 solution in water. 3. Mix this solution with acetic acid in a 2:7 ratio. Thorough mixing requires ∼ 20–30 min on the magnetic stirring plate. The resulting solution has a brown color. For further information about storage and preparation see Note 10. A slightly different, simpler version of reducing solution is described in Note 11. A recently proposed ascorbic acid/acetic acid reducing solution which has been shown to be specific for nitrite is described in Note 12. The simplicity and specificity for nitrite could soon make this new method very popular in clinical practice. 2.3.2. Chemiluminescence – Instrumentation

Currently, there are two commercially available NO analyzers: Sievers NO analyzer (NOA, model 280i, GE Analytical Instruments, Boulder, CO, USA) and CLD 88Y (EcoPhysics, Duernten, Switzerland). The Sievers NOA requires an O2 tank attached to the analyzer as a source of ozone (O3 ), while the CLD 88Y derives O3 from O2 in normal atmospheric conditions. Inert gas (He, N2 , or Ar) from a source tank with a

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constant flow rate of 100–150 mL/min is used as the NO carrier in both models. Sensitivity of both analyzers for NO detection is comparable and is in the low nM/high picomolar range of nitrite. According to manufacturers, it is possible to detect upward from 1 pM of NO gas in the reaction chamber. However, the sensitivity of each individual unit should be determined using calibration either with a known amount of NO gas or using NO donors.

3. Methods 3.1. Nitrite-Preserving “Stop” Solution

3.1.1. Stop solution is added to blood sample in a ratio 1:4 (v/v, stop solution/sample). Sample is then vortexed vigorously for ∼10 s to allow complete lysis of cells and thorough mixing. Samples containing Hb will become brown, which indicates the presence of oxidized heme in the form of metHb. 3.1.2. Stop solution will cause lysis of cells; so if it is desirable to measure nitrite content in cells and supernatant separately, intact cells must be separated from supernatant prior to addition of stop solution (Fig. 4.1).

3.2. Sample Collection and Pretreatment 3.2.1. Whole Blood Treatment

Collected blood samples are aliquoted into 1.5–2 mL Eppendorf tubes containing stop solution in a 1:4 ratio (v/v, stop solution/sample). For preparation of nitrite preserving stop solution, see previous Section 2.1. After vigorous vortexing, samples can be frozen on dry ice and stored at –80◦ C for later analysis (stable up to several months). If processed immediately, the deproteination step follows.

3.2.2. RBC Treatment

Whole blood samples are centrifuged at 5,000×g for 5 min to separate RBC from plasma. Collected RBCs are aliquoted into 1.5–2 mL tubes containing stop solution in a 1:4 ratio (v/v, stop solution/RBC). After vigorous vortexing, samples are either frozen on dry ice and stored at –80◦ C for delayed processing or measured immediately after deproteination (see Section 3.2.4).

3.2.3. Plasma Treatment

Whole blood samples are centrifuged at 5,000×g for 5 min to separate RBC from plasma. Collected supernatant – plasma – is aliquoted into 1.5–2 mL tubes. Stop solution is not needed if plasma is clear and without hemolysis. If significant hemolysis is

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observed, stop solution must be used as described above for RBC treatment. Aliquots can be either frozen on dry ice and stored at –80◦ C for delayed processing or measured immediately. Deproteination is not needed for plasma. 3.2.4. Deproteination Step

Whole blood and RBC aliquots must be deproteinated before injection into the chemiluminescence reaction vessel to avoid excessive foaming. If processing previously frozen samples, they should slowly thaw on ice. Proteins are precipitated by adding cold methanol to samples in a 1:1 ratio (v/v) and mixture vortexing. Proteins are then removed from mixture by centrifugation of the mixture at 13,000×g for 5 min at 4◦ C. Supernatant is carefully transferred to a new 1.5 mL tube and used for nitrite analysis (see Notes 13 and 14). It is necessary to keep small aliquots (∼1 mL) of all solutions used in your sample treatments––stop solution, methanol, water, or any additional treatment that was used. All these solutions might contain nitrite contamination, which will contribute to the nitrite concentration measured in samples. Nitrite content in all these solutions is determined at the same time as nitrite in samples and, if necessary, corrections for nitrite contamination from these external sources can be made.

3.3. Chemiluminescence– Data Acquisition

There is a slight difference in initial set up of the CLD 88Y and Sievers NOA instruments, but data collection and processing are similar. CLD 88Y is slightly more user-friendly than Sievers NOA.

3.3.1. Procedure for CLD 88Y from EcoPhysics

1. Turn on the main switch located on the rear panel. 2. Turn on the computer attached to the instrument via the USB cable and start the CLD Excel macro. 3. The computer should detect the connected instrument and start running CLD macro, displaying a signal from the photomultiplier in millivolts as a function of time (see Note 15). 4. Check that input gas line connector located on the rear panel is tightly screwed on and that there is a dry HPLC filter between the bubbler (filled with 12 mL of 1 M sodium hydroxide (NaOH) to completely cover glass frit) and instrument. Any water residues left in the line or filter will result in an unstable baseline. Do not connect the line between bubbler and instrument at this point. 5. Wait for the CLD 88Y to automatically turn on the vacuum pump to evacuate the instrument’s internal reaction chamber. This will show on the computer screen as a large decrease in signal. Wait until an appropriate vacuum in the

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chamber is achieved and the baseline is stable – the complete process takes ∼1–1.5 h. 6. Connect the carrier gas input line from the gas tank to the appropriate port of the reaction vessel. Start the carrier gas flow prior to pouring I3 solution into the glass vessel. Carrier gas should flow at a rate that allows the addition of 7–9 mL of I3 solution into the reaction vessel without letting the liquid go below the glass frit. Adjust the gas flow rate so that I3 solution is gently bubbling but not overflowing beyond the cap or into the cooling part of reaction vessel. 7. Add 50–100 μL of Antifoam B into I3 solution in the reaction vessel (see Note 16). 8. Connect the NaOH-filled bubbler to the input gas line with HPLC filter. The second port of the bubbler should be still open with gentle bubbling observed, as the CLD 88Y creates a slight negative pressure to aspirate the gas from the line. If bubbling is missing, check all connections, including those on the HPLC filter, for tightness and all lines for some mechanical obstruction or humidity. 9. Carefully connect the remaining gas output line from the reaction vessel to the bubbler, watching for any possible I3 solution overflow that can be prevented by adjusting carrier gas flow. Once the carrier gas flow is adjusted, it has to remain the same for the duration of the experiment. If it is necessary to adjust the flow during the experiment, a new standard curve needs to be acquired at this flow rate. A detailed diagram of the lines and reaction vessel connections is given in Fig. 4.3. 10. About ∼30 min after all lines are connected the baseline should be flat and relative noise should not exceed 0.1–0.3 mV. Possible reasons for a high/unstable baseline and ideas for troubleshooting are in Note 17. 11. Stop the acquisition and run a new CLD Excel macro file. After acquiring a few minutes of a steady baseline, proceed with nitrite standards and sample measurements. 3.3.2. Procedure for Sievers NOA Setup Is Described Separately in Note 18

While waiting for the NO analyzer to warm up and stabilize, wash all Hamilton syringes you plan to use thoroughly. See Note 19 for precautions and useful tips regarding syringes. Prepare 1 μM sodium nitrite standard solution (NaNO2 , MW 68.99) for the calibration curve. After the NO analyzer is ready for measurements, acquire the chemiluminescence signal from your samples and nitrite standards as follows: 1. Using a well-washed Hamilton syringe, inject 50 μL of 1 μM nitrite solution into I3 reaction mixture. Wait between injections until the chemiluminescence signal peak drops

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back to baseline and baseline is stable for at least 2 min to achieve good separation of peaks. Repeat injection of 50 μL two more times to get triplicates of each data point. Wash syringe after each injection as described in Note 19. 2. To acquire nitrite standards for the calibration curve, inject in triplicates of 100, 150, 200, 250, and 300 μL of 1 μM nitrite solution. In each case, take care to wait until the signal drops back to the baseline and then acquire 1–2 min of baseline. The calibration curve and its uses are described in Note 20. 3. Nitrite standards are usually measured before samples. If it later becomes apparent that nitrite levels of some measured samples are out of the range of injected standards (peak heights differ considerably), a second set of nitrite standards with appropriate amounts of nitrite should be injected after all samples have been measured. This set of injections is then used as the calibration curve. 4. If nitrite levels and other NOx combined levels (R-SNO + RNNO + nitrosylHb (HbNO)) are expected to be comparable, pretreatment with acid sulfanilamide (AS, MW 172.21) and separate measurement of total NOx and nitrite-depleted sample is necessary – for AS treatment procedure and for general guidelines of data acquisition. We recommend use of AS treatment on the first trial for all samples; once the ratio of NOx /nitrite is determined to be reasonably low, neglecting the NOx contribution and skipping AS treatment is acceptable (see Note 9 and Fig. 4.2). 5. At a minimum, triplicates of all samples must be measured. Injection amounts may vary from 50 μL up to 300–500 μL, depending on expected nitrite content. All peaks must be clearly distinguished from baseline/noise variation. Multiple injections of 500 μL are not recommended, as this will quickly fill up the reaction vessel. 6. Each time an aliquot is injected into the reaction vessel; record the time of injection, what was injected and in what volume. If any dilutions or additional processing of the samples occurred, notes should be made to permit accurate tracking of all dilutions of the original sample. 3.4. Chemiluminescence – Data Processing

After all samples and nitrite standards are measured, nitrite levels in injected aliquots and original samples can be determined. We import the data into Origin software (OriginLab Corp. Northampton, MA, USA), but any other software capable of determining the area under peaks can be used Excel (Microsoft), Sigmaplot (Systat Software), Prizm (GraphPad Software) or similar.

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Data acquired using CLD88Y are saved in an Excel file. This file does not contain any information about the origin of injection therefore separate records must be kept manually. Sievers NOA uses its proprietary Liquid software to acquire data into two separate files. “Filename.data” contains data and “filename.info” with time of mark and comments. Both formats, Excel and “filename.data” are standard; most spreadsheet-based software will open them directly. To process acquired data (Fig. 4.2). 1. Open data file in Origin (or other software). 2. Determine the area under the peaks using baseline markers placed at the peak take off and after the signal returns to baseline again. 3. If treatment with AS is used, correct the areas under the peaks for NOx contribution accordingly (see Note 9 and Fig. 4.2). 4. Using the calibration curve, determine the amount of nitrite in each aliquot injected into the reaction vessel, N (see Note 20). 5. Determine the total dilution factor of original sample in the aliquot, D. For example, if the sample was diluted in a 1:9 ratio (v/v, sample/buffer), D=1/10=0.1. 6. Calculate the fraction of the volume of original sample, VS that was present in the total volume of injected aliquot, VT . 7. Divide the amount of nitrite in aliquot N (determined in point 3) by the volume of the original sample: N/VS or N/(D/VT ). This is the concentration of nitrite in the original sample.

4. Notes 1. The reaction of nitrite with heme-containing proteins represents the major route for nitrite destruction in any biological sample. This is especially important in blood, as hemoglobin is present in millimolar concentrations. The reaction of nitrite with ferrous hemoglobin, in its oxy- or deoxy- state, leads to oxidation of nitrite to nitrate and ferrous heme to ferric heme and, in the case of deoxyHb to a new reaction product, nitrosylHb (HbNO). Ferric heme in metHb does not react further with nitrite. Nitrite has a low affinity for ferric heme and the metHb–nitrite complex formation is accepted. However, this weakly bound complex is formed only in excess of nitrite and its formation will not

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affect results when chemiluminescence is used to measure nitrite levels. Summary of reactions of oxyHb and deoxyHb with nitrite (reactions 1 and 2) and NO (reactions 3 and 4): oxyHb + NO2 – → metHb + NO3 – (reaction 1) deoxyHb + NO2 – → metHb + NO (reaction 2) deoxyHb + NO → HbNO (reaction 3) oxyHb + NO → metHb + NO3 – (reaction 4) 2. “Stop” solution is a mixture of the following reagents: K3 Fe(CN)6 , NEM, NP-40, and water. K3 Fe(CN)6 is the main “nitrite-preserving” ingredient. The product of its reaction with oxyHb and deoxyHb is metHb. NEM is a thiol reactive compound commonly used to modify cysteine residues in proteins and peptides; here it is used to prevent additional R-SNO formation that would increase the signal detected by I3 -based chemiluminescence. NP40 is a nonionic detergent used to assure complete lysis of membranes and fast mixing of K3 Fe(CN)6 and NEM with the entire contents of the cell. Triton X-100 can be used instead of NP-40. 3. Only water with low-nitrite content, such as Milli-Q treated water or molecular biology grade water, should be used for preparation of samples and solutions, as well as for final wash of syringes and all glassware. 4. Stop solution should be prepared daily by mixing 1.32 g K3 Fe(CN)6 and 0.0665 g NEM in 4.5 mL water, with addition of 0.5 mL NP-40. The solution should be kept at 4◦ C and can be stored for a few days. However, for best results it is recommended to use fresh solution for sample preparation, whenever possible. 5. Modified stop solution: KCN, K3 Fe(CN)6 , NEM, NP-40, H2 O. In this modification reported in (2), KCN (potassium cyanide, MW 65.12, highly toxic) is added to the stop solution mixture at a final concentration of 200 mM and dissolved together with K3 Fe(CN)6 and NEM before final mixing with NP-40. Addition of KCN leads to formation of cyanometheme and cyanomethemoglobin(MetHbCN), which further prevents oxidized heme from forming a complex with free nitrite. From our experience, it is not necessary to use KCN for blood samples, but we recommend it for processing tissue samples, as many active heme proteins are in the ferric oxidation state.

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6. All necessary materials and stop solution should be prepared in advance and kept at hand – immediate access to a tabletop centrifuge cooled to 4◦ C and a vortex are strongly suggested if nitrite measurement in RBC’s are required. Once blood is collected, it is critical to work as fast as possible, as nitrite decays in the blood with half-life of ∼11– 13 min (6). 7. Principle of chemiluminescence-based detection of NO: NO + O3 → NO2 ∗ + O2 (reaction 5) NO2 ∗ → NO2 + hv (photon)(reaction 6) 8. In tri-iodide (I3 – ) solution, nitrite is reduced to NO by its reaction with iodide ion as follows: NO2 – + H+ ⇔ HNO2 (reaction 7) 2I– + 2HNO2 + 2H+ → 2NO + I2 + 2H2 O (reaction 8) 9. I3 -based reducing solution is not selective for nitrite. NO will be released also from R-SNO, R-NNO, and Fe-NO functional groups. Nitrate will not be reduced in I3 solution. The “additional” or “false” nitrite signal can be accounted for by using nitrite-specific sample treatments with AS. Nitrite reacts with sulfanilamide in an acidic environment and irreversibly forms diazonium cation complex that is not reduced to NO in I3 solution (4, 6). It is also possible to separate and quantify contributions from R-SNO and Fe-NO using different targeted sample pretreatments (1–4, 7). Procedure to separate the nitrite-related chemiluminescence signal from total NOx signal using AS treatment: a. 5% solution of acidified sulfanilamide (5% wt/v in 1 M HCl or 290 mM solution of sulfanilamide) is prepared by dissolving 0.5 g of sulfanilamide in 10 mL of 1 M HCl. b. Divide the sample into two aliquots, A and B. c. Aliquot A is mixed well with AS solution in a ratio 1/9 (v/v, AS/sample) and incubated for 3 min to deplete free nitrite. NOx level is measured using the standard method. d. Water or buffer is added into aliquot B at the same ratio as above to account for dilution, and NOx levels are measured using the standard method. e. True nitrite signal is calculated by subtracting AS-treated sample A from non-treated sample B as shown in Fig. 4.2.

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f. At normal physiological conditions nitrite is present in blood in at least 10-fold excess over other NOx metabolites (excluding nitrate), therefore the AS treatment is necessary only if highly precise determination of nitrite concentration is required. We recommend first running a few pilot aliquots with and without AS treatment to estimate the contribution of other NOx metabolites and decide accordingly. 10. Best results are obtained with freshly prepared I3 solution used within 1 day and kept preferably in a dark bottle, as I-containing reagents are light sensitive. We prepared 90 mL of I3 solution by dissolving 1 g KI and 0.65 g I2 in 20 mL water with subsequent addition of 70 mL acetic acid. This amount is enough to refill the reaction vessel ∼10 times. 11. Nitrite can be reduced to NO using a mixture of KI or NaI (sodium iodide, MW 149.89, light sensitive) and glacial acetic acid by the same mechanism as I3 solution. This original mixture was thought to be specific for nitrite. However, it has been shown that the I3 – ion, which can also reduce R-SNO and R-NNO, forms in the reaction vessel over time and the real selectivity toward nitrite can be only achieved in freshly prepared KI/acetic acid solutions and is maintained only for the first few sample injections. These requirements make the use of KI/acetic acid solution impractical, and it is therefore currently not as widespread as the I3 -based method (3). a. Reactants: KI, glacial acetic acid, H2 O. NaI can be used instead of KI. b. Prepare 667.15 mM solution of KI in H2 O. Mix this solution with glacial acetic acid in 1:9 proportions. This reaction mixture is dark yellow and is light sensitive. It can be kept in dark bottle at room temperature and used within 1 day. An easy way to prepare a sufficient supply for 1 day (50 mL) is to dissolve 553.7 mg of KI (or 500 mg of NaI) in 5 mL of H2 O, then add 45 mL of acetic acid. 12. A mixture of ascorbic acid (MW 176.12, not stable in solution, prepare fresh every day) and acetic acid has been recently proposed for selective reduction of nitrite in plasma for chemiluminescence detection (5). This method is specific for nitrite; the reaction mixture will not release NO from any other NOx metabolites. Nitrite is reduced to NO using a mixture of ascorbic acid and glacial acetic acid by disproportionation as shown in reactions 9–13. Reactions 9 and 10 will occur in glacial acetic acid alone, while in the presence of ascorbic acid, acting as a reducing

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agent, the reaction equilibrium shifts toward NO release, as shown in reactions 11–13. NO2 – + H+ ⇔ HNO2 (reaction 9) HNO2 + H+ ⇔ H2 NO2 + ⇔ NO+ + H2 O (reaction 10) HNO2 + Asc → NO + H2 O + Asc• (reaction 11) H2 NO2 + + Asc →NO + Asc• + H2 O + H+ (reaction 12) NO+ + Asc → NO + Asc• + H+ (reaction 13) a. Reactants: ascorbic acid, glacial acetic acid, and H2 O. b. Prepare 500 mM ascorbic acid in water. Mix this solution with glacial acetic acid in a ratio 1:7 to prepare the reaction mixture. c. According to (5), completion of nitrite reduction depends on ascorbic acid concentration; and at least 50 mM ascorbic acid is recommended for complete reduction of plasma nitrite. However, we believe that few pilot experiments with different concentrations of ascorbic acid and nitrite standards in the expected nitrite concentration range are necessary prior to final sample measurements. 13. For deproteination, some authors use methanol in a ratio 1:1 (v/v) with multiple centrifugations at 750×g for 2 min each until the supernatant appears clear (5) or in a ratio of 1:2 (v/v, sample/methanol) with a single centrifugation at 21,000×g at 4◦ C for 15 min (8). 14. Deproteination step significantly dilutes the nitrite in samples, and it is not recommended when extremely low nitrite levels are expected. In such a case, we recommend the use of a twofold increase in antifoam and direct injection of samples containing protein. After each injection, frequent changes of the reaction mixture will be required. 15. If an error message “connection not found” is displayed and the Excel macro does not run, check the USB connector/replace cable and restart the computer. If the problem persists, the driver for the communication port needs to be reinstalled. 16. Biological samples containing large amounts of protein must be either completely deproteinated prior to injection into reaction vessel, or antifoaming agent needs to be added to the I3 mixture. Often samples are only partially deproteinated so there is still a need for antifoam. We use antifoam B emulsion with 50–100 μL of antifoam for 9 mL of reaction solution, added directly into reaction vessel (1). Others recommend antifoam 204 organic (3) or antifoam SE-15 (8). It is important to remember that antifoam itself

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may contain some nitrite impurities, so it is recommended to wait after antifoam addition until all nitrite is consumed and the baseline is stable again. 17. High baseline signals occur for several reasons: humidity – water droplets or dust in the gas line or filter, high NOx pollution in air, loose connections between reaction vessels and the instrument, cap of the reaction vessel not fully closed, or an old Teflon septum. From all listed, the most frequent reasons are loose connectors and an old Teflon septum. 18. Sievers NOA instrument set up: All glassware is connected to Sievers NOA the same way as described for CLD 88Y in Section 3.3.1 and shown in Fig. 4.3. The acquired data are processed as described in Sections 3.3 and 3.4. a. Open O2 tank connected to the instrument. NOA derives O3 from the attached O2 tank and will give an error message if O2 supply is inadequate or missing. b. Choose option “analysis” from the main menu on the instrument panel by highlighting it and push “enter.” On the next screen choose “start” and press “enter.” The vacuum pump should start at this point. Wait for cooler of the photomultiplier to reach a temperature below –12◦ C and the vacuum in the reaction chamber to reach approximately 6 Torr. The baseline should stabilize itself below 50 mV – the absolute value depends on the starting state of the instrument and it takes between 30 min and several hours. The baseline has to be stable within 1–2 mV, the nominal value for the baseline is less important than its stability, as all measurements are relative to the baseline. However, the lower the initial baseline and the lower the cooler temperature will give more precision and more sensitivity. c. When temperature and pressure are stable, turn on the computer attached to the instrument and start Labview-based Liquid software. To achieve communication between the instrument and acquisition software, data output from the instrument has to be enabled as follows: press “enter” when in “data menu” – screen will read “output disabled” – press “enter” to change it to “enabled” and then press “clear” to return back to data screen. If preparation of a new sample is done while the instrument is running, data transfer should be disabled when a measurement is not taken – see point 7.b. below for a detailed explanation. d. Wait for a stable baseline and start injecting samples and nitrite standards into I3 reaction solution as described

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already for CLD 88Y. Always wait for a stable baseline after the peak. The Liquid software has an option to “mark” times of injection and it will export them into a separate file (filename.info) together with any written comments – this is a useful complement to the data file (filename.data). e. Once all samples and standards are measured, choose the “stop” option in the Liquid software menu – this will write data from a temporary file into a permanent file known as “filename.data” (see also point 7.b) f. To stop NOA analyzer, disconnect the He tank from the glass reaction vessel first, then press “clear” to get into main menu. Choose “analysis” option, then “stop” and confirm “stop” on the confirmation screen. Turn off supply of O2 from O2 tank. g. NOA troubleshooting tips: • Instrument should be always left in standby mode, unless physically moved to another location. After turning it off for a longer period of time (1–2 days), a few days of constant running are needed to stabilize the baseline again. • It is possible to enable/disable the data output as many times as necessary during one session of Liquid software. The software will display an error message when communication is disabled, but it will continue running and writing into the same file after communication is enabled again. This feature is important because Liquid software runs only for a preset amount of time (maximum continuous run time is 1 h) and will discard all data unless “stop” button is chosen before the run time expires. • If the “error” message in Liquid software persists after communication was enabled, check if the baud rate for the communication port, set for the instrument, and in the software match. To check baud rate on the instrument: from main menu, choose option “control,” and then “setup.” There are two options in this menu: “view” and “change.” Choosing “view” only shows the current setup, “change” will allow modifications. Choosing either of them will display the next menu with the option “configuration” and then “COM port.” This option displays the current baud rate and changes are made using “up” and “down” arrows keys. From the software side, baud rate is setup from the first pop-up window after the software starts.

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19. We use Hamilton syringes (size 100–500 μL) with sharp needle tips to inject small aliquots of samples into I3 solution through the Teflon septum of the reaction vessel cap. For the best results use gastight syringes; however, good results are also obtained with ordinary Hamilton syringes that are new and still tight. It is necessary to thoroughly wash each syringe prior to use, as well as before and after sample injection. High-quality nitrite-free water, the same grade that is used to prepare all solutions and samples, is also used to wash syringes. We use two 50 mL Falcon centrifuge tubes filled with Milli-Q treated water to wash our syringes and control their cleanness as follows: use the first Falcon tube to wash the syringe by repeating a fill up/empty cycle at least 15–20 times. After cleaning, fill the syringe with 100 μL of water from the second tube and inject the water into I3 solution in the reaction vessel. If the running baseline of chemiluminescence signal changes, i.e., a peak appears, repeat the washing procedure until no peak or only a very small peak of residual nitrite is observed. In case of any residual peak, multiple water injections permit later estimation of nitrite contamination in the water. If the “water peak” persists, use a new bottle of water (verify that Millipore system measures conductivity of ∼18 M cm, if not – change cartridges) and/or replace the Hamilton syringe. It is absolutely crucial for determination of precise nitrite levels to minimize nitrite contamination from external sources. 20. Determination of a calibration curve and its use: a. Measure the area under each peak for all nitrite standards. b. Plot these areas as a function of total amount of nitrite for each injection. Total amount of nitrite in 50, 100, 150, 200, 250, and 300 μL of 1 μM nitrite solution is 50, 100, 150, 200, 250, and 300 pM. Recalculate the amount of nitrite injected if different volumes were used. c. Calculate the slope, K, of the standard curve. d. To determine the total amount of nitrite in the sample, divide this area by the slope (total amount of nitrite in sample) = (area under peak)/K. 21. Additional information can be found in references (9–13). References 1. Pelletier, M. M., Kleinbongard, P., Ringwood, L., Hito, R., Hunter, C. J., Schechter, A. N., et al. (2006) The mea-

surement of blood and plasma nitrite by chemiluminescence: pitfalls and solutions. Free Radic Biol Med 41, 541–548.

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2. Yang, B. K., Vivas, E. X., Reiter, C. D., Gladwin, M. T. (2003) Methodologies for the sensitive and specific measurement of S-nitrosothiols, iron-nitrosyls and Nitrite in biological samples. Free Radic Res 37, 1–10. 3. Pinder, A. G., Rogers, S. C., Khalatbari, A., Ingram, T. E., James, P. E. (2008) The measurement of nitric oxide and its metabolites in biological samples by ozone-based chemiluminescence.. In (J.T. Hancock ed.), Methods in Molecular Biology, Redox-Mediated Signal Transduction, vol. 476. Humana press, Totowa, NJ, pp. 11–28. 4. Mac Arthur, P. H., Shiva, S., Gladwin, M. T. (2007) Measurement of circulating nitrite and S-nitrosothiols by reductive chemiluminescence. J Chromatogr B 851, 93–105. 5. Nagababu, E., Rifkind, J. M. (2007) Measurement of plasma nitrite by chemiluminescence without interference of S-, N-nitroso and nitrated species. Free Radic Biol Med 42, 1146–1154. 6. Tsikas, D. (2007) Analysis of nitrite and nitrate in biological fluids by assay based on Griess reaction: appraisal of the Griess reaction in the L-arginine/nitric oxide area of research. J Chromatogr B 851, 51–70. 7. Bryan, N. S., Grisham, M. B. (2007) Methods to detect nitric oxide and its metabolites in biological samples. Free Radic Biol Med 43, 645–657.

8. Hendgen-Cotta, U., Grau, M., Rasaaf, T., Gharinin, P., Kelm, M., Kleinbongard, P. (2008) Reductive gas-phase chemiluminescence and flow injection analysis for measurement of nitric oxide pool in biological matrices. Method Enzymol 441, 295–315. 9. Wang, X., Bryan, N. S., MacArthur, P. H., Rodriguez, J., Gladwin, M. T., Feelisch, M. (2006) Measurement of nitric oxide levels in the red cell. J Biol Chem 281, 26994–27002. 10. Dejam, A., Kleinbongard, P., Rasaaf, T., Hamada, S., Gharni, P., Rodriguez, J., Feelisch, M., Kelm, M. (2003) Thiols enhance NO formation from nitrate photolysis. Free Radic Biol Med 35, 1551–1559. 11. Laver, J. R., Stevanin, T. M., Read, R. C. (2008) Chemiluminescence quantification of NO and its derivatives in liquid samples. Method Enzymol 436, 113–127. 12. Rasaaf, T., Feelisch, M., Kelm, M. (2004) Circulating NO pool: assessment of nitrite and nitroso species in blood and tissues. Free Radic Biol Med 36, 413–422. 13. Cornelius, J., Tran, T., Turner, N., Piazza, A., Mills, L., Slack, R., Hauser, S., Alexander, J. S., Grisham, M., Feelisch, M., Rodriguez, J. (2009) Isotope tracing enhancement of chemiluminescence assays for nitric oxide research. Biol Chem 390, 181–189.

Chapter 5 Selective Fluorescent Activation for Bioimaging the Expression of Nitric Oxide in Cellular and In Vivo Systems Junfeng Zhang and Hao Hong Abstract Nitric oxide serves as a messenger for cellular signaling and physiological reactions such as inflammatory responses in vivo. Fluorescent bioimaging can be a useful tool in nitric oxide functional research. However, current nitric oxide in vivo imaging protocols often result in suboptimal image quality. Selective fluorescent activation of probes after their reaction with nitric oxide is an appropriate way for imaging nitric oxide. Fluorescent naphtho[2,3-d]imidazol derivatives can react with copper II to form nonfluorescent compounds, which can then enable direct imaging of nitric oxide in vitro or in vivo, based on the redox action of copper II. These probes can be applied to image nitric oxide produced by inducible nitric oxide synthase in cellular systems such as lipopolysaccharide-activated murine macrophages. More importantly, the probes can be utilized for in vivo imaging of nitric oxide production, for example, in acute severe hepatic injury. This chapter describes the methods required to apply such probes as nitric oxide bioimaging agents with potential use for diagnostic and pathological studies of nitric oxide-related diseases. Key words: Fluorescent bioimaging, nitric oxide, in vivo.

1. Introduction Nitric oxide (NO), an uncharged free radical, is an intracellular and intercellular messenger that governs a myriad of physiological and pathophysiological processes. Since its initial discovery as an endothelium-derived relaxing factor in 1987 (1, 2), it has been well established that NO is a ubiquitous messenger in many biological events such as vascular homeostasis, neurotransmission, immune systems, and tumor progression (3–5). NO is a doubleedged sword in a number of diseases, especially cancer (5–9). The H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_5, © Springer Science+Business Media, LLC 2011

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exact function of NO depends on its temporal and spatial distribution. Much research has been carried out to unveil the true nature of NO in order to cure or alleviate related diseases. However, NO is highly reactive and can be converted to other species very quickly; thus, it is very difficult to detect and monitor the level of NO in vivo. As an important tool in cell- and small-animal-based research, fluorescent imaging provides many unique capabilities which derives from its intrinsic molecular sensitivity, resolution span, repeatability, high level of safety, and relatively low instrumentation costs (10). There are some successful small-moleculebased fluorescent probes for NO, which have widely used biological applications, including o-diaminofluoresceins (DAFs) (11), diaminorhodamines (DARs) (12), and o-diaminocyanines (DACs) (13). However, the fluorescent alterations of those probes react with oxidized NO products rather than NO itself, which limits their practical use in certain environments (e.g., imaging of NO in an anoxic cell culture or an anoxic tissue such as a tumor). Moreover, NO has a half-life of up to 7.5 min under certain circumstances (14), so the fluorescent imaging of its oxidized products cannot accurately represent real-time NO production. The Massachusetts Institute of Technology has designed novel compounds (15, 16) to react directly with NO. These compounds can serve as real-time cellular fluorescent detectors for NO; however, considering their molecular mobility and stability, no in vivo detection results have been reported so far. Fluorescent imaging of NO using small-molecule-based probes with selective activation for NO provides a distinct in vivo imaging capability. In this chapter, we will introduce 2 hydroxyphenyl-1H-naphtho[2,3-d]imidazol compounds, a series of highly fluorescent molecules, which can form stable nonfluorescent reactive compounds with Cu(II). After their reaction with Cu(II), the formed products will serve as useful inducible probes for visualizing NO in vitro or in vivo. Two compounds were selected: 4-methoxy-2-(1H-naphtho[2,3-d]imidazol-2-yl)phenol (MNIP) and 5 -chloro-2-(2 -hydroxyphenyl)-1H-naphtho [2,3-d]imidazol (CHNI) to react with Cu(II) to form MNIP-Cu and CHNI-Cu. These compounds are specifically designed to detect inflammatory NO in vivo.

2. Materials 2.1. Compounds for Synthesis of MNIP or CHNI

1. 4-Methoxy-salicylaldehyde. 2. 5-Chloro-salicylaldehyde. 3. Naphthalene-1,2-diamine.

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4. Methanol, ethanol, dimethylformamide (DMF), acetic acid, n-hexane, acetonitrile, ammonium acetate, dimethyl sulfoxide (DMSO), and ethyl acetate. 5. Mn(II)Cl2 ·4H2 O. 2.2. Cell Culture

1. Dulbecco’s Modified Eagle’s Medium (DMEM) and fetal bovine serum (FBS); DMEM is supplemented with 10% FBS. 2. Solution of trypsin (0.25%) and ethylenediamine tetraacetic acid (EDTA) (1 mM). 3. Lipopolysaccharide (LPS) was dissolved in 1× phosphate buffered saline (PBS) to a stock solution of 10 μg/mL stocked separately in 1 mL volume at –20◦ C. 4. H2 O used in cell culture was all purified in a Milli-Q water purification system.

2.3. RT-PCR

1. TriPure isolation reagent kit (total RNA extraction kit). 2. Oligo (dT)15 primers, AMV reverse transcriptase, dNTPs. 3. iNOS (+) primer: 5 -CTGCAGCACTTGGATCAGGAACC TG-3 , iNOS (–) primer sequence: 5 -GGGAGTAGCC TGTGTGCACCTGGAA-3 , GAPDH (+) primer sequence: 5 -AACGACCCCTTCATTGACC-3 , GAPDH (–) primer sequence: 5 -TCAGATGCCTGCTTCACC-3 . 4. Agarose and 10× TAE electrophoresis buffer.

2.4. Hepatic Injury Model

1. D-galactosamine. 2. Male Balb/c mice (6–8 weeks, 18–20 g). 3. Gadolinium chloride (GdCl3 ). 4. Test kit for NO (Merck, Darmstadt, Germany). 5. Optimum cutting temperature (OCT) compound (Sakura Finetek Co. Tokyo, Japan).

3. Methods The fluorescent ligand MNIP or CHNI is synthesized using a manganese-assisting method. Manganese triacetate is rapidly evolving as a new and exceptionally versatile reagent in organic synthesis. Recently, it has been widely employed in the synthesis of many kinds of heterocyclic compounds (17). We did a one-pot synthesis of MNIP or CHNI producing a high yield by the oxidative condensation of 4-methoxy-salicylaldehyde for MNIP or 5chloro-salicylaldehyde for CHNI with naphthalene-1,2-diamine

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using manganese triacetate as a relatively benign oxidizing reagent under milder reaction conditions. Synthesized MNIP/CHNI can react with Cu(II) to form the nonfluorescent reaction compounds MNIP-Cu or CHNI-Cu. These compounds can react directly, rapidly, and specifically with NO over other interfering reactive nitrogen species (RNS) or reactive oxygen species (ROS) to activate fluorescence. These cell-permeable coordination compounds can image NO produced in LPS-activated Raw264.7 cells with good resolution. In an acute severe hepatic injury (ASHI) model using Balb/c mouse induced by LPS and GalN (18), we delivered MNIP-Cu/CHNI-Cu via intravenous injection, and found a significant increase in fluorescence in the liver and blood. To further explore NO fluctuation in the liver, we undertook injections through the liver portal vein with MNIP-Cu/CHNI-Cu in normal or ASHI mice. Fluorescent images of liver sections indicated that these compounds could directly image NO generation in vivo in a manner directly proportional to immunological insult. 3.1. Preparation of CHNI and MNIP, CHNI-Cu, and MNIP-Cu

1. For CHNI, dissolve 30 mM 5-chloro-salicylaldehyde (for MNIP, it is 30 mM 4-methoxy-salicylaldehyde) in 125 mL methanol by continuous stirring. 2. Add 15 mM naphthalene-1,2-diamine to the solution. 3. Stir at room temperature for 1 h or until appropriate sediments are formed. 4. Collect sediments via filtration and wash with methanol. 5. Vacuum dry the washed sediments to acquire the intermediate ligands. 6. 13 mM of the intermediate ligand was dissolved in 100 mL chloroform to form solution 1. 7. 2.5 g Mn(II)Cl2 ·4H2 O was dissolved in 40 mL ethanol to form solution 2. 8. Combine solutions 1 and 2 forming a brown-colored solution 3. 9. Stir solution 3 for 24 h with bubbled air at room temperature. After 12 h a dark brown solution will form. After 24 h dark brown sediments should form. 10. Collect the sediments via filtration and wash with DMF followed by an ethanol wash. 11. Vacuum dry the sediment to obtain the manganese (III) complex. 12. Dissolve 200 mg of manganese (III) complex in 15 mL of 33% methanol and adjust to pH 10 using NaOH.

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13. Stir the solution for 12 h at room temperature and remove the formed sediments by filtration. 14. Adjust the filtrate to pH 5 using acetic acid (CH3 COOH). 15. Elute the filtrate using ethyl acetate. 16. Dry the eluted filtrate using anhydrous sodium sulfate to form CHNI or MNIP. 17. Purify CHNI or MNIP by thin-layer chromatography where ethyl acetate/n-hexane (1:1) serves as the mobile phase. 18. Recrystallize the purified chemicals using ethanol to obtain pure CHNI or MNIP. 19. Dissolve CHNI or MNIP in DMSO to form a solution with a concentration of 1 mM. 20. Add 50 μM copper sulfate (CuSO4 ) to the pure CHNI or MNIP solution to achieve a final molar ratio of CHNI/MNIP: copper = 1:1 (see Note 1). 21. Mix at room temperature for 30 min to form a stable yellow solution of CHNI-Cu/MNIP-Cu. The whole synthesis scheme is shown in Fig. 5.1.

Fig. 5.1. a The synthesis scheme of MNIP and MNIP-Cu. b The synthesis scheme of CHNI and CHNI-Cu.

3.2. Characterization of CHNI, MNIP, CHNI-Cu, and MNIP-Cu

1. CHNI and MNIP were characterized by 1 H NMR, 13 C NMR (DRX-500, Bruker, Switzerland), EI-MS (JMS-700, JOEL, Japan), and elemental analysis (JSX-3202 M, JOEL, Japan). NMR spectra were recorded in d6-DMSO with tetramethylsilane (TMS) serving as the internal standard. 2. CHNI was obtained as light yellow microcrystals, 177 mg (60%), mp = 360◦ C (from ethanol); 1 H NMR (400 MHz,

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deuteriodimethyl sulfoxide): δ = 13.33 (br s, 2H, OH and NH), 8.27 (d, 1H, arom H, J = 2.8 Hz), 8.20 (br s, 2H, arom H), 8.05 (m, 2H, arom H), 7.48 (dd, 1H, arom H, J = 2.8, 8.8 Hz), 7.43 (m, 2H, arom H), 7.12 (d, 1H, arom H, J = 8.8 Hz); 13 C NMR (100 MHz, deuteriodimethyl sulfoxide): δ = 157.3, 154.3, 132.02, 132.0, 130.21, 130.20, 127.70, 127.66, 127.60, 126.12, 126.07, 123.88, 123.84, 122.7, 119.1, 119.0, 113.5; EI-MS: m/z (relative intensity) = 296 (34, M+ +2), 294 (100, M+ ), 260 (10), 231 (10), 149 (12), 140 (16), 115 (20), 57 (12). Anal. Calcd. for C17 H11 ClN2 O: C, 69.28; H, 3.76; N, 9.50. Found: C, 68.98; H, 3.83; N, 9.25. 3. MNIP was obtained as yellow needles, 253 mg (58%), mp 299–300◦ C (from ethanol); 1 H NMR (400 MHz, deuteriodimethyl sulfoxide) δ 13.50 (br s, 1H, NH), 13.03 (br s, 1H, OH), 8.18 (br s, 1H, arom H), 8.05 (d, 2H, arom H, J = 8.8 Hz), 8.04 (br s, 1H, arom H), 8.02 (m, 2H, arom H), 7.40 (m, 2H, arom H), 6.68 (dd, 1H, arom H, J = 2.8, 8.8 Hz), 6.63 (d, 1H, arom H, J = 2.8 Hz), 3.84 (s, 3H, OCH3 ); 13 C NMR (100 MHz, deuteriodimethyl sulfoxide) δ = 162.8, 162.6, 156.0, 141.6, 141.5, 130.0, 127.98, 127.94, 127.85, 127.82, 123.56, 123.50, 113.67, 113.62, 106.7, 105.2, 101.4, 55.4; EI-MS, m/z (relative intensity) 290 (100, M+ ), 261 (120), 247 (23), 219 (6), 193 (4). Anal. Calcd. for C18 H14 N2 O2 : C, 74.47; H, 4.86; N, 9.65. Found: C, 74.21; H, 4.95; N, 9.48. 4. CHNI-Cu and MNIP-Cu were characterized by MALDITOF (Ultraflex II, Bruker, Switzerland), ESR (FE1X, JOEL, Japan), and FT-IR (Nicolet Nexus 870, Thermal Fisher Scientific, USA). The representative MALDI-TOF, FT-IR, and ESR spectra of CHNI-Cu are shown in Fig. 5.2. 3.3. Reaction and Fluorescence Alteration of CHNI-Cu/MNIP-Cu with NO 3.3.1. Formation of NO and NO Standard Solution

1. Degas all apparatus with N2 for 30 min to exclude O2 as NO is rapidly destroyed by O2 . 2. Slowly drop 2 M H2 SO4 into a saturated NaNO2 solution to generate NO (19). 3. Bubble 10 mL of deoxygenated distilled H2 O for 30 min and maintain under NO atmosphere until use to produce a saturated NO solution (at 20◦ C, NO ≈ 1.8 mM (20)) (see Note 2). NO gas is toxic at concentrations higher than 100 ppm, so the bubbling procedure was carried out in a

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Fig. 5.2. Characterization of CHNI, CHNI-Cu, and the reaction products of CHNI-Cu with NO. a MALDI-TOF of CHNI-Cu and its reaction product with NO. b FT-IR of CHNI, CHNI-Cu, and its reaction product with NO. c Reaction mechanism specification of CHNI-Cu with NO. d ESR spectra of CHNI-Cu and its reaction product with NO. e The fluorescence enhancement curve of CHNI-Cu with different amount of NO.

fume hood. Standard solutions were freshly prepared for each experiment and kept in a glass flask with a rubber septum. Dilutions of the saturated solution were made using deoxygenated H2 O samples. 3.3.2. Fluorescence Assay with NO (CHNI-Cu as the Example)

The fluorescence measurements were carried out on an F-4500 model spectrophotometer (Hitachi, Tokyo, Japan). A slit width of 1 nm was used for both excitation and emission with a photomultiplier voltage of 700 V. Absorption and emission spectra were recorded in 10% DMSO/90% double distilled H2 O, consistent with all the following cellular experiments. The testing concentration was 1 μM. At 0, 0.5, 1, and 5 min after the reaction with 1,000 equivalent of NO (see Note 3), the fluorescence curves were recorded. The fluorescence alteration curve of CHNI-Cu with NO was recorded in Fig. 5.3.

3.3.3. The Reaction Mechanism with NO (Part I: CHNI-Cu)

Two hours after the reaction of CHNI-Cu with 1,000 equivalent of NO (see Note 3), the products were analyzed by FT-IR and MALDI-TOF. At the same time, ESR spectra were used to monitor the Cu(II) alteration between CHNI-Cu and its reaction

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Fig. 5.3. Fluorescence curve and specific test of CHNI-Cu after reaction with NO.

product with NO (see Note 4). The ESR, FT-IR, and MALDITOF spectra are shown in Fig. 5.2 with the specified reaction mechanism. During the reaction of CHNI-Cu, NO oxidizes the Cu(II) to form N-nitrosated fluorescent CHNI-NO ligand. 3.3.4. The Reaction Mechanism with NO (Part II: HPLC, MNIP-Cu)

The HPLC column used was C18, 4.6 × 250 mm, 5 μm, 300 Å. The mobile phase, acetonitrile (60% v/v), 0.01 M ammonium acetate, and 0.5% CH3 COOH (40%), was pumped with a flow rate of 0.5 mL/min. The injection volume for each sample was 20 μL. The detection system was Shimadzu SPD-10A (Shimadzu, Japan) with a UV detection wavelength of 360 nm and a sample concentration of 250 μM. To confirm the elution time of MNIP-NO, 250 μM MNIP-Cu and NO were reacted and analyzed. To confirm that the application of MNIP-Cu in LPS-activated Raw264.7 cells generated the fluorescent MNIP-NO, Raw264.7 cells were stimulated with 500 ng/mL LPS. MNIP-Cu was added

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simultaneously to a final concentration of 50 μM. Cells (5 × 106 ) were digested and lysed in 5 mL PBS after 12 h. 1 mL of chloroform was added to the cell lysate to extract the fluorescent products. A solution of 50 μM MNIP-Cu was also extracted with chloroform to serve as a nonreacting control. After the chloroform was vaporized, the samples were resolved in 100 μL acetonitrile before analysis. The elution time of MNIP was 1.982 min. In MNIP-Cu solution, three new elution peaks appeared at 2.365 min (CUSO4 ), 4.482 min (MNIP:Cu2+ = 1:1), and 5.615 min (MNIP:Cu2+ = 2:1). Judging from the integral areas, the structure of MNIP:Cu2+ = 1:1 is the main product. The reaction of MNIP-Cu with NO produced a peak at 3.948 min which indicated that this was the reaction product of MNIP-Cu with NO (MNIP-NO). Meanwhile, when MNIPCu was extracted with chloroform, only MNIP and MNIP:Cu2+ = 1:1 compounds were extracted in the chloroform, with no evidence of an MNIP:Cu2+ = 2:1 compound existing in the chloroform extraction. Similar to the chemical environments, after MNIP-Cu was applied to LPS-stimulated Raw264.7 cells, the emergence of a peak at 3.948 min in the cell extract further proved that MNIP-Cu reacted with cell-released NO to generate the N-nitrosated fluorescent product MNIP-NO. 3.3.5. Specificity Test for NO (CHNI-Cu)

Fluorescence responses of CHNI-Cu were recorded after the − + addition of 100 equivalent of H2 O2 , NO− 2 , NO3 , NH4 , − ONOO , LPS, GalN, NO, or equal volume of FBS for 2 h in DMSO/water (v/v = 1:9) solution. The excitation wavelength was 353 nm. The fluorescence intensities are normalized with respect to that of CHNI-Cu. The testing concentration of CHNICu is 1 μM. Specific test results are shown in Fig. 5.3.

3.4. Application of MNIP-Cu In Vitro 3.4.1. In Vitro Treatment with MNIP-Cu ± LPS

The MNIP-Cu solution was sterile filtered through a 0.22 μm filter. Raw264.7 cells were grown in DMEM supplemented with 10% FBS to a confluence of 85%. The cells were incubated with FBS-supplemented DMEM and treated with 500 ng/mL LPS. MNIP-Cu was added to the culture media at a final concentration of 10 μM in 10% DMSO/FBS-supplemented DMEM (see Note 5). The fluorescence images were recorded at 2 h intervals using a fluorescent microscope. At each time point, cells were rinsed twice with 1× PBS and illuminated with a high-pressure mercury lamp (330–385 nm band-pass filter) and observed using an emission filter (BA420). Untreated control Raw264.7 cells were incubated with MNIP-Cu to evaluate baseline fluorescence in quiescent cells. Cellular fluorescent images are recorded in Fig. 5.4.

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Fig. 5.4. Application of MNIP-Cu in macrophages. a Images at different time points of LPS stimulation. The final concentration of MNIP-Cu is 10 μM. 1,400 W and PTIO serves as negative controls. RT-PCR was used to monitor the iNOS expression at according time point. b Semiquantification of fluorescence intensity at different time points.

3.4.2. In Vitro NO Deprivation

To confirm cellular fluorescence was associated only with NO production, we used the following reagents: an inducible nitric oxide synthase (iNOS) inhibitor, N-[3-(aminomethyl) benzyl]acetamidine, dihydrochloride (1,400 W) (21), an NO scavenger, 2-phenyl-4,4,5,5-tetramethylimidazoline-3-oxide-1oxyl (PTIO) (see Note 6). All LPS treatments were supplemented with BHT (100 μM), SOD (100 U/mL), and catalase (100 U/mL). 1,400 W treatment group: Pretreat the cells to 100 μM 1,400 W for 4 h. Then add 500 ng/mL LPS and 10 μM MNIP-Cu. PTIO treatment group: Expose the cells to 500 ng/mL LPS. Posttreat with 500 μM PTIO added at 3, 5, 7, 9, 11, 13, or 19 h after LPS. Thirty minutes after each treatment with PTIO, add 10 μM MNIP-Cu. Fluorescence images were recorded at 4, 6, 8, 10, 12, 14, and 20 h after LPS addition. Images from various time points were analyzed using ImageJ software (US National Institutes of Health; version 1.38×). Fluorescence intensity per cell area at each time point was calculated using a minimum of five images.

3.4.3. Measurement of iNOS Expression In Vitro Using RT-PCR

Total RNA was extracted from cells using the TriPure isolation reagent kit. The yield and quality of the RNA were assessed

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by measuring absorbance at 260 and 280 nm followed by electrophoresis on 1% agarose gel. 1 μg of RNA was reversetranscribed into cDNA by using oligo (dT)15 primers and AMV reverse-transcriptase at 42◦ C for 1 h. Sequence-specific primers for iNOS and the housekeeping gene GAPDH were used for cDNA amplification. Denature cDNA at 95◦ C for 5 min with thermal cycling conditions as follows: 95◦ C for 25 s; 55◦ C for 45 s, 72◦ C for 30 s, with a final extension step at 72◦ C for 10 min. PCR products were electrophoresed through 1% agarose gel, stained with ethidium bromide, and visualized under ultraviolet illumination. 3.5. Establishment of In Vivo Models

All the animal studies complied with the current ethical regulations for animal research at Nanjing University. Fifty Balb/c mice were randomly divided into two groups (20 mice in the control group and 30 mice in the treatment group) (see Note 7).

3.5.1. Establishment of ASHI In Vitro Model

NaCl (0.9%, 0.2 mL) was injected intraperitoneally in the control group. LPS (5 μg/kg) together with GalN (400 mg/kg) in 0.9% NaCl was injected intraperitoneally in the test group. All animals were fasted 12 h before respective treatments.

3.5.2. Depletion of Kupffer Cells in the ASHI Model

Macrophages, especially Kupffer cells, were depleted in vivo 24 h after i.v. injections of gadolinium(III) chloride (GdCl3 ). Intravenous injections of GdCl3 (10 mg/kg) were administered to deplete Kupffer cells at 24 h before LPS/GalN induction of ASHI.

3.6. In Vivo Application of CHNI-Cu in ASHI Mice 3.6.1. Fluorescence Enhancement Evaluation of CHNI-Cu in the Main Organs of Normal/ASHI Mice

Six hours after ASHI induction, CHNI-Cu (10 mg/kg, dissolved in 0.2 mL of 50% DMSO and 50% NaCl) was delivered via i.v. tail vein injection. At 2 h post i.v. injection, mice were sacrificed and the heart, liver, spleen, lung, kidney, brain, and blood were taken out and homogenized in 400 μL DMSO. The homogenates were centrifuged at 1,500 rcf for 10 min. 150 μL of the supernatants were removed for fluorescent assay using a Safire fluorescence reader (Tecan Co., Zurich, Switzerland). Figure 5.5a shows the organ fluorescence intensity enhancement in ASHI mice compared to normal mice. From the figure, the fluorescence intensities in liver, spleen, and blood are very high compared to other organs in ASHI mice. To normalize for total protein content, the specific organ protein levels were determined by colorimetric assay using Coomassie brilliant blue G-250.

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Fig. 5.5. Application of CHNI-Cu in ASHI mice. a Fluorescence enhancement in the main organs of ASHI mice compared to those of normal mice after CHNI-Cu application. All the data were normalized with respect to the fluorescent intensities in the organs of control mice. Error bars indicate SD. b NO content in the main organs of ASHI mice and normal mice acquired by NO test kit to serve as a comparison. Error bars represent SD.

3.6.2. Evaluation of NO with CHNI-Cu in the Main Organs of Normal/ASHI Mice

The test kit for NO was used for analyzing the NO contents in the main organs and blood, which serve as a comparative model (Fig. 5.5b). In this assay, the main organs were homogenized in 400 μL 0.9% NaCl solution. The homogenates were centrifuged at 1,500 rcf for 10 min. 150 μL of the supernatants were taken for NO quantification via the protocol listed in NO test kit. Comparatively similar NO expression levels and CHNI-Cu fluorescence were observed in the liver, spleen, and blood.

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Six hours after respective treatment with LPS/GalN, 10 mg/kg CHNI-Cu was injected into the hepatic portal vein. 0.5 h postinjection, mice were sacrificed and the livers were excised and soaked in an optimum cutting temperature compound (Sakura Finetek Co., Tokyo, Japan) for 1 h at −22◦ C. These treated tissues were cut into 4 μm sections using a Sakura coldtome model cm-41 (Tokyo, Japan). Acetone at 4◦ C was added to the sections to fix the tissues. One hour postfixation, the sections were imaged via fluorescence microscopy (IX71 fluorescent microscope). Representative images are shown in Fig. 5.6.

Fig. 5.6. Bioimaging of NO using CHNI-Cu in in vivo inflammatory system.

3.6.4. Imaging of NO in Kupffer Cell-Depleted Liver

Twenty-four hours before the LPS/GalN treatment, i.v. injections of GdCl3 (10 mg/kg) were administered (see Note 8). 10 mg/kg CHNI-Cu was injected into the hepatic portal vein 6 h after LPS/GalN treatment. Mice were sacrificed 0.5 h postinjection and the liver excised for sectioning. The macrophagedepleted sections were imaged via fluorescence microscopy (IX71 fluorescent microscope) (see Note 9). Representative images are shown in Fig. 5.6.

4. Notes 1. During the formation of CHNI-Cu/MNIP-Cu, CUSO4 solution should be dropped slowly into the CHNI (MNIP) solution with continuous stirring. An excess of cupric ions

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should be added if the formed solution is to be stored for longer periods. 2. Unless stated otherwise, all the aqua solutions should be prepared with H2 O which has a resistivity of no less than 18.2 M cm. 3. The NO standard solution should be used within 2 h and the dilution of the solution to different concentrations should be carried out in deoxygenated H2 O. 4. To reveal the true reaction mechanism between CHNI-Cu and NO, the ESR or NMR measurement should be undertaken immediately or within 2 h of the reaction. N2 or Ar protection of the test tube should be adopted to prevent copper oxidization. 5. The concentration of CHNI-Cu in vitro should be no more than 100 μM (for MNIP, 150 μM), otherwise CHNI-Cu can form fluorescent crystals intracellularly which may interfere with the assay. 10 μM is an optimal concentration for imaging of NO in most cells. 6. This protocol has a high sensitivity for other in vitro systems and can detect nM fluctuations of NO. Immunocells, vascular cells (e.g., HUVEC cells (22)), and tumor cells are three examples in which these probes have broad application. 7. Establishing the effects of the ASHI model in different strains can be variable. With our experience, Balb/c and C57BL/6 mice are two appropriate strains for the ASHI model. Although the imaging effects may be better in some C57BL/6 mice, the results are more stable in Balb/c mice. 8. Effective depletion of Kupffer cells is dependent upon quality and quantity of GdCl3 . GdCl3 from Sigma-Aldrich has the least toxicity and the best depletion effects. A dose range between 5 and 15 mg/kg are applicable for introducing depletion of Kupffer cells. 9. The distribution of CHNI-Cu/MNIP-Cu in the liver is generally uniform. Thirty minutes postinjection is adequate for the generation of high-quality fluorescent images. Increasing postinjection time did not enhance imaging. References 1. Ignarro, L. J., Buga, G. M., Wood, K. S., Byrns, R. E., Chaudhuri, G. (1987) Endothelium-derived relaxing factor produced and released from artery and vein is nitric oxide. Proc Natl Acad Sci 84, 9265–9269. 2. Palmer, R. M., Ferrige, A. G., Moncada, S. (1987) Nitric oxide release accounts for the

biological activity of endothelium-derived relaxing factor. Nature 327, 524–526. 3. Garthwaite, J. (2008) Concepts of neural nitric oxide-mediated transmission. Eur J Neurosci 27, 2783–2802. 4. Bogdan, C. (2001) Nitric oxide and the immune response. Nat Immunol 2, 907–916.

Expression of Nitric Oxide in Cellular and In Vivo Systems 5. Fukumura, D., Kashiwagi, S., Jain, R. K. (2006) The role of nitric oxide in tumour progression. Nat Rev Cancer 6, 521–534. 6. Sullivan, R., Graham, C. H. (2008) Chemosensitization of cancer by nitric oxide. Curr Pharm Des 14, 1113–1123. 7. Mocellin, S., Bronte, V., Nitti, D. (2007) Nitric oxide, a double edged sword in cancer biology: searching for therapeutic opportunities. Med Res Rev 27, 317–352. 8. Ridnour, L. A., Thomas, D. D., Donzelli, S., Espey, M. G., Roberts, D. D., Wink, D. A., Isenberg, J. S. (2006) The biphasic nature of nitric oxide responses in tumor biology. Antioxid Redox Signal 8, 1329–1337. 9. Hirst, D., Robson, T. (2007) Targeting nitric oxide for cancer therapy. J Pharm Pharmacol 59, 3–13. 10. Moriyama, E. H., Zheng, G., Wilson, B. C. (2008) Optical molecular imaging: from single cell to patient. Clin Pharmacol Ther 84, 267–271. 11. Kojima, H., Nakatsubo, N., Kikuchi, K., Kawahara, S., Kirino, Y., Nagoshi, H., Hirata, Y., Nagano, T. (1998) Detection and imaging of nitric oxide with novel fluorescent indicators: diaminofluoresceins. Anal Chem 70, 2446–2453. 12. Kojima, H., Hirotani, M., Nakatsubo, N., Kikuchi, K., Urano, Y., Higuchi, T., Hirata, Y., Nagano, T. (2001) Bioimaging of nitric oxide with fluorescent indicators based on the rhodamine chromophore. Anal Chem 73, 1967–1973. 13. Sasaki, E., Kojima, H., Nishimatsu, H., Urano, Y., Kikuchi, K., Hirata, Y., Nagano, T. (2005) Highly sensitive near-infrared fluorescent probes for nitric oxide and their application to isolated organs. J Am Chem Soc 127, 3684–3685. 14. Nagano, T., Yoshimura, T. (2002) Bioimaging of nitric oxide. Chem Rev 102, 1235–1270.

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15. Lim, M. H., Xu, D., Lippard, S. J. (2006) Visualization of nitric oxide in living cells by a copper-based fluorescent probe. Nat Chem Biol 2, 375–380. 16. Lim, M. H., Lippard, S. J. (2005) Copper complexes for fluorescence-based NO detection in aqueous solution. J Am Chem Soc 127, 12170–12171. 17. Ouyang, J., Hong, H., Zhao, Y., Shen, H., Shen, C., Zhang, C., Zhang, J. (2008) Bioimaging nitric oxide in activated macrophages in vitro and hepatic inflammation in vivo based on a coppernaphthoimidazol coordination compound. Nitric Oxide 19, 42–49. 18. Oberholzer, A., Oberholzer, C., Bahjat, F. R., Edwards, C. K., Moldawer, L. L. (2001) Genetic determinants of lipopolysaccharide and D-galactosamine-mediated hepatocellular apoptosis and lethality. J Endotoxin Res 7, 375–380. 19. Huang, K. J., Wang, H., Ma, M., Zhang, X., Zhang, H. S. (2007) Real-time imaging of nitric oxide production in living cells with 1,3,5,7-tetramethyl-2,6-dicarbethoxy8-(3 ,4 -diaminophenyl)-difluoroboradiazas-indacence by invert fluorescence microscope. Nitric Oxide 16, 36–43. 20. Hong, H., Sun, J., Cai, W. (2009) Multimodality imaging of nitric oxide and nitric oxide synthases. Free Radic Biol Med 47, 684–698. 21. Garvey, E. P., Oplinger, J. A., Furfine, E. S., Kiff, R. J., Laszlo, F., Whittle, B. J., Knowles, R. G. (1997) 1400 W is a slow, tight binding, and highly selective inhibitor of inducible nitric-oxide synthase in vitro and in vivo. J Biol Chem 272, 4959–4963. 22. Ouyang, J., Hong, H., Shen, C., Zhao, Y., Ouyang, C., Dong, L., Zhu, J., Guo, Z., Zeng, K., Chen, J., Zhang, C., Zhang, J. (2008) A novel fluorescent probe for the detection of nitric oxide in vitro and in vivo. Free Radic Biol Med 45, 1426–1436.

Chapter 6 Real-Time Measurement of Murine Hippocampus NO Levels in Response to Cerebral Ischemia/Reperfusion Xiaoxiang Zheng, Kezhou Liu, and Yong Yang Abstract Nitric oxide has been implicated as a mediator of synaptic transmission and a pathological factor in stroke/reperfusion. The purpose of this study was to detect the change of nitric oxide concentration in rat hippocampus during global cerebral ischemia and reperfusion in vivo and to reveal effects of different nitric oxide synthases. In the present study, the real-time record of nitric oxide levels in rat hippocampus was obtained by using a nitric oxide sensor during global cerebral ischemia and the initial stage of reperfusion. We also observed the effects of two inhibitors of nitric oxide synthases on nitric oxide concentration. The two inhibitors were administrated intravenously at the onset of reperfusion and 1 h later. The change of the nitric oxide concentration in the initial stage of reperfusion was 0.768 ± 0.029 μM. 7-Nitroindazole (7-NI, inhibitor of nNOS) had a strong inhibitive effect on nitric oxide synthesis at both time points, while 1400 W dihydrochloride (1400 W, inhibitor of iNOS) had no significant effect on nitric oxide synthesis. The results showed that during the initial stage of reperfusion, nitric oxide biosynthesis was mainly an nNOS-dependent process. Key words: Nitric oxide, stroke/reperfusion, in vivo, iNOS, nNOS.

1. Introduction Following the discovery that the endothelium-derived relaxing factor (EDRF) and nitric oxide (NO) are identical (1, 2), there have been an abundance of studies investigating the physiological and pathological roles of NO. As an important molecular messenger, NO functions as a nerve mediator, as well as a detrimental factor in the nervous system. In central and peripheral nervous systems, NO participates in many forms of synaptic transmission and pathological progress (3). The dual function of this molecule H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_6, © Springer Science+Business Media, LLC 2011

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is of particular interest during cerebral ischemia and reperfusion (I/R). The role of NO pathogenesis in ischemic brain damage is controversial (4, 5). Evidence suggests that NO can play both neurotoxic and neuroprotective roles during the process of cerebral ischemia and reperfusion (2, 3). As a result of the multiple and contradictory actions of NO as well as the different sources of nitric oxide synthase (NOS), modulations of NO in ischemic brain have been shown to produce a variety of outcomes (6). It is thought that neuronal NOS (nNOS) and inducible NOS (iNOS) are detrimental to the ischemic brain, whereas endothelial NOS (eNOS) activity may be protective (7–9). However, relatively little information is available on the specific contributions of the three NOS isoforms with relation to the ischemic brain, e.g., precise enzyme activity and corresponding yield of NO production. As the effect of NO is variable depending upon the stage of global ischemia, a comprehensive understanding of the real-time synthesis of the NOS isoforms is required. A number of techniques have been described to measure NOS enzymatic activity indirectly by quantifying the production of nitrite and nitrate, or directly by measuring NO levels in tissues using spin trapping and electron paramagnetic resonance spectroscopy (10). Therefore, continuous and real-time measurement of NO concentration would be beneficial with regard to determining the effect of NO in global cerebral ischemia and reperfusion (11). Neishi et al. (12) used a catheter-type NO sensor for measurement of NO in a canine coronary sinus and concluded that this method was useful for evaluating the bioavailability of NO in coronary circulation (12). In the present study, we measured NO levels in rat hippocampus using a carbon-fiber NO sensor during the process of global cerebral ischemia and the initial stage of reperfusion. The CA1 area of the rat hippocampus has already been proven to express copious levels of the various NOS enzymes (13, 14). Meanwhile, we separately observed the effects of two different NOS inhibitors on the production of NO in the process of global cerebral ischemia. Using these direct measurement techniques, we were able to untangle the complex interactions between cerebral global ischemia and NO expression.

2. Materials 2.1. NO Sensor

The ISO-NOPF200 NO probe is manufactured by World Precision Instruments (WPI), Sarasota, USA. The lowest detection limit of this probe is 0.2 nM and the linear range is 10 nM–1 μM. Previous experiments were already done to identify the sensitivity and selectivity of the probe (15) (see Note 1). The diameter

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of the probe is 200 μm, and the diffusion of the NO molecule to the surface of the carbon fiber takes 3 s (16), which is shorter than the half-life of NO. Before each experiment, polarize the electrode (see Note 2) and calibrate with S-nitroso-N-acetyl - DLpenicillamine (SNAP) (see Notes 3, 4, 5, and 6). The conversion efficiency of SNAP to NO is 100% (17). Regression analysis has previously demonstrated that the probe has an excellent linearity (r 2 ≥ 0.99) for NO measurement (Fig. 6.1).

Fig. 6.1. A calibration curve of the NO sensor. Linear regression analysis showed the NO sensor had a satisfactory linearity (r 2 = 0.99923). The relationship of NO concentration (Y ), equal to the SNAP concentration (see Note 6), and the voltage output of recorder (X ) was Y = 13,200 ∗ X + 28.33.

2.2. Laser Doppler Flowmetry

Monitor the cerebral blood flow (CBF) by Laser Doppler Flowmetry (LDF) (PeriFlux5000, Perimed, Stockholm, Sweden) during the global cerebral ischemia and reperfusion. Drill a burr hole (1 mm in diameter) 5.2 mm posterior and 4.2 mm lateral to the Bregma without injury to the dura mater. Place the laser Doppler probe (Probe407-1 Perimed, Stockholm, Sweden) in the burr hole. Measure CBF continuously (τ = 0.3) from before the onset of global ischemia until 30 min after reperfusion. The results of Laser Doppler Flowmetry should indicate that global cerebral ischemia and reperfusion has occurred. Typical results of the Laser Doppler Flowmetry are shown in Fig. 6.2.

2.3. Drugs

The narcotics, chloral hydrate and urethane, and the inhibitors of NOS, 7-nitroindazole (7-NI), dihydrochloride (1400 W), aminoguanidine (AG) and SNAP were used in this experiment. AG was noted to have a strong influence on the NO sensor (data not shown); therefore, 7-NI and 1400 W should be preferentially selected as inhibitors of nNOS and iNOS.

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Fig. 6.2. Dynamic changes in CBF measured by LDF. Arrows indicate clamping both carotid arteries and removal of the nontraumatic artery clamps.

3. Methods 3.1. NO Measurement

3.2. Animal Preparation

Insert the NO probe into the CA1 area of the rat hippocampus by stereotaxic apparatus (see Note 7). According to the stereotaxic atlas of rat brain, the fixed position of the sensor should be 5.2 mm posterior, 4.2 mm lateral, and 7.0 mm ventral, relative to the Bregma (controlateral to the position of the laser Doppler probe), which is the central part of the hippocampus and close to the cranial bone. At this position, the Ag/AgCl reference electrode of the probe is also immerged in the brain. After each experiment, remove the brain and immerse in 10% formalin saline for 8 h. Frozen transverse sections (2 mm) should be stained with cresyl violet to identify the electrode tracks. Using the ISO-NO recorder, the output signal of the probe can be determined and recorded with a sampling frequency of 100 Hz (see Note 5). The signal is then transformed from current to voltage by the recording software (Chart 5.0, ADI, USA). In the calibration experiment, a standard calibration curve should be constructed according to the recorded data, which should reveal linear relationship between NO concentration and the output voltage. Then, according to this calibration curve, the change of the NO concentration during reperfusion can be calculated by the change of the voltage output (see Note 8). 1. Anesthetize rats intraperitoneally with 10% chloral hydrate (4 mL/kg). 2. Electrocauterize the two vertebral arteries in the bony tunnel of the first vertebra according to the method of Pulsinelli W.A. (18), using an electrocoagulator-dessicator (see Notes 9 and 10).

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3. After 24 h, anesthetize the same rat intraperitoneally with 20% urethane (7 mL/kg). 4. Make a low cervical incision to expose the common carotid arteries. Isolate but do not occlude both carotid arteries with a 4.0 silk tie, ensuring preservation of the vagus nerve. 5. Completely restrict blood supply by clamping both carotid arteries for 20 min using nontraumatic artery clamps. Recirculate blood flow by releasing the clips to recover blood supply. 6. Throughout the experiment, maintain the body temperature at 37±1◦ C (see Note 4). 3.3. Experimental Protocol

1. The procedures of experimentation included separation of the two common carotid arteries by cotton thread, cannulation of the left carotid vein for drug injection, placement of a laser Doppler probe for CBF detection, and insertion of the NO probe into the CA1 area of hippocampus by stereotaxic apparatus. 2. After the NO probe is inserted into the hippocampus, it should be allowed to stabilize for about 1 h until the basal line is uniform (see Note 7). 3. Global cerebral ischemia is then induced by occlusion of both carotid arteries, and recirculation of blood flow established by releasing the nontraumatic artery clamps. According to our previous experiments, long-time global cerebral ischemia resulted in serious damage to the brain. Thus we carefully chose the occlusion time as 20 min since this amount of time is enough to cause brain damage to release NO, and also guarantee the rats’ survival during the experiment. The changes of NO concentration and the standard error from the mean for NO concentrations at various time points are shown in Fig. 6.3. 4. At the moment of onset of reperfusion and 1 h later, 7-NI (50 mg/kg) and 1400 W (20 mg/kg) should be separately injected into the left carotid vein. The current responses of the electrode to NO generation are electronically recorded. Figure 6.4 shows previous results.

4. Notes 1. NO diffuses through a selective membrane covering the NO sensor and is oxidized at the working electrode, resulting in an electrical (redox) current. This redox current is

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Fig. 6.3. Changes in NO concentration in rat hippocampus during global ischemia/reperfusion. a The statistical results of NO concentration profiles during brain global ischemia/reperfusion (n = 6). b The mean NO concentration at various time points: (a) 1 min before global ischemia (representing the normal physiological condition), (b) 10 min after global ischemia (the minimum value in global ischemia), (c) 20 min after global ischemia (the platform period value in global ischemia), (d) 15 min after reperfusion (the summit value in perfusion), and (e) 40 min after reperfusion (the platform period in reperfusion). ∗ , p < 0.01 vs. the normal physiological condition, #, p < 0.01 vs. the summit of reperfusion.

proportional to the concentration of NO. The NO sensor can be used to detect NO in vivo. 2. Before each calibration and experiment, the NO sensor must be connected to the recorder for 4–5 h in order to be polarized. 3. Accurate measurements of NO using the ISO-NO require an accurate calibration. Our calibration method is based on the decomposition of the SNAP. Calibration must be performed before each experiment. The calibration range should be kept close to the anticipated experimental concentration of NO.

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Fig. 6.4. The effect of 7-NI and 1400 W to NO concentration in the rat hippocampus during global ischemia/reperfusion. a 7-NI was administrated at the onset of reperfusion. b 7-NI was administrated 1 h later after reperfusion. c 1400 W was administrated at the onset of reperfusion. d 1400 W was administrated 1 h after reperfusion.

4. The NO sensor is sensitive to temperature, and large temperature changes can cause changes to the baseline. Therefore it is recommended that any calibration is performed at the same temperature as the experiment. 5. The ISO-NO system must be properly grounded with the experimental animal, not only during experiment but also in the calibration. 6. Standard SNAP solution is a key point in the calibration. According to the manual, dissolve 5 mg EDTA in 250 mL of HPLC water and then add 5.6 mg SNAP to the solution. The decomposition of SNAP at low temperatures in the dark and in the absence of trace metal ions proceeds very slowly due to the presence of the chelating reagent, EDTA. Since the SNAP solution is very sensitive to light and temperature, it should be stored in the dark and in a refrigerator until required. 7. It takes about 1 h for the NO sensor to be stable when immersing its tip in a new solution or biological tissue. 8. The NO sensor can continuously work for about 8 h in our experiment. For longer experiments, fatigue of the sensor should be considered. 9. The global cerebral ischemia model is performed according to reference (18). Using stereotaxic apparatus may help expose the vertebral arteries more easily.

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10. After the cauterization of the vertebral arteries, the rat needs 24 h to recover and adapt to the reduced blood supply to the brain provided by the two carotid arteries.

Acknowledgement This work was supported by Zhejiang Provincial Key Laboratory of Chinese Medicine Screening, Exploitation & Medicinal Effectiveness Appraise for Cardio-Cerebral Vascular & Nervous System. References 1. Palmer, R. M., Ferrige, A. G., Moncada, S. (1987) Nitric oxide release accounts for the biological activity of endothelium-derived relaxing factor. Nature 327, 524–526. 2. Ignarro, L. J., Buga, G. M., Wood, K. S., Byrns, R. E., Chaudhuri, G. (1987) Endothelium-derived relaxing factor produced and released from artery and vein is nitric oxide. PNAS 84, 9265–9269. 3. Moncada, S., Bolanos, J. P. (2006) Nitric oxide, cell bioenergetics and neurodegeneration. J Neurochem 97, 1676–1689. 4. Choi, D. W. (1993) Nitric oxide: foe or friend to the injured brain? PNAS 90, 9741–9743. 5. Dawson, T. M., Snyder, S. H. (1994) Gases as biological messengers: nitric oxide and carbon monoxide in the brain. J Neurosci 14, 5147–5159. 6. Panahian, N., Yoshida, T., Huang, P., Hedley-Whyte, E., Dalkara, T., Fishman, M., Moskowitz, M. (1996) Attenuated hippocampal damage after global cerebral global ischemia in mice mutant in neuronal nitric oxide synthase. Neuroscience 72, 343–354. 7. Iadecola, C. (1997) Bright and dark sides of nitric oxide in ischemic brain injury. Trends Neurosci 20, 132–139. 8. Samdani, A. F., Dawson, T. M., Dawson, V. L. (1997) Nitric oxide synthase in models of focal global ischemia. Stroke 28, 1283–1288. 9. Adachi, N., Lei, B., Soutani, M., Arai, T. (2000) Different roles of neuronal and endothelial nitric oxide synthases on ischemic nitric oxide production in gerbil striatum. Neurosci Lett 288, 151–154. 10. Khoo, J., Alp, N., Bendall, J., Kawashima, S., Yokoyama, M., Zhang, Y., Casadei, B., Channon, K. (2004) EPR quantification of vascular nitric oxide production in geneti-

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16. 17.

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cally modified mouse models. Nitric Oxide 10, 156–161. Shin, J., Schoenfisch, M. (2006) Improving the biocompatibility of in vivo sensors. Analyst 131, 609–615. Neishi, Y., Mochizuki, S., Miyasaka, T., Kawamoto, T., Kume, T., Sukmawan, R., Tsukiji, M., Ogasawara, Y., Kajiya, F., Akasaka, T., Yoshida, K., Goto, M. (2005) Evaluation of bioavailability of nitric oxide in coronary circulation by direct measurement of plasma nitric oxide concentration. PNAS 102, 11456–11461. Valtschanoff, J. G., Weinberg, R. J., Kharazia, V. N., Nakane, M., Schmidt, H. H. H. W. (1993) Neurons in rat hippocampus that synthesize nitric oxide. J Comp Neurol 331, 111–121. Bredt, D. S., Hwang, P. M., Snyder, S. H. (1990) Localization of nitric oxide synthase indicating a neural role for nitric oxide. Nature 347, 768–770. Liu, K., Ning, G., Zheng, X. (2005) In vivo detection of nitric oxide in rat hippocampus. In Proceedings of the 2005 IEEE Eng Med Biol 27th Annual Conference, September 1– 4, 2005, Shanghai, China, pp. 1039–1042. Zhang, X., Cardosa, L., Broderik, M., Hein, H. (1999) An integrated nitric oxide selective ultramicrosensor. Free Radic Biol Med 27, 89. Zhang, X., Cardosa, L., Broderick, M., Fein, H., Davies, I. R. (2000) Novel calibration method for nitric oxide microsensors by stoichiometrical generation of nitric oxide from snap. Electroanalysis 12, 425–428. Pulsinelli, W. A., Buchan, A. M. (1988) The four-vessel occlusion rat model: method for complete occlusion of vertebral arteries and control of collateral circulation. Stroke 19, 913–914.

Chapter 7 Detection of Low Levels of Nitric Oxide Using an Electrochemical Sensor Yong Chool Boo, Gyeong In Mun, Sarah L. Tressel, and Hanjoong Jo Abstract Nitric oxide produced from nitric oxide synthases mediates various physiological and pathological events in biological systems. However, quantitative assessment of nitric oxide from biological sources remains a difficult task. Here we describe a procedure for the quantification of low levels of nitric oxide using a nitric oxide – selective electrochemical sensor. Nitric oxide is oxidized to nitrite and/or nitrate and accumulated in the aqueous media. First, nitrate in biological fluids or culture media is converted to nitrite by an enzymatic method. Nitrite is then chemically converted to equimolar NO in an acidic iodide bath, where nitric oxide is detected by the sensor. Using this method, the present study demonstrates siRNA-mediated suppression of nitric oxide synthase 3 leading to a significant decline of basal nitric oxide production in human umbilical vein endothelial cells. Basal nitric oxide production from HUVECs is also shown to be inhibited by NG -nitro-L-arginine methyl ester but not by NG -nitro-D-arginine methyl ester (D-NAME). The analytical method presented here provides a sensitive and convenient tool for measuring basal and stimulated nitric oxide production from biological sources. Key words: Nitric oxide, nitrate, nitrite, electrochemical sensor.

1. Introduction The biological importance of nitric oxide (NO) has been well established in the past decades. As a consequence, various methods have been developed to directly or indirectly assess the low levels of NO produced from biological sources. They include spectrophotometric, fluorometric, electrochemical, chemiluminescence, and electron spin resonance methods. Indeed, some H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_7, © Springer Science+Business Media, LLC 2011

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of these methods appear to be highly sensitive and selective, but they also require expensive and highly technical equipment making them inaccessible to many potential users. NO is very unstable with a half-life of 2–30 s in aqueous media and rapidly reacts with molecular oxygen to form nitrite (1). In the presence of oxidizing species such as oxyhemoproteins, nitrite is further oxidized to nitrate (2). Nitrite or nitrate (collectively referred to as NOx) accumulate in extracellular fluids such as sera, urine, and cell culture media (3). Therefore the detection of NOx has been a useful method to assess NO production from biological sources (4, 5) when direct monitoring of NO is impractical. Berkels et al. (6) reported a method to measure NOx by converting them to NO which is detected by an amperometric NO sensor (6). This method is attractive because it provides a rapid and easy assay while requiring relatively inexpensive instruments that are affordable for laboratories. Furthermore, a new version of the NO sensor with higher sensitivity has become commercially available (7). Here we present a detailed procedure for the quantification of NOx using a new version of the NO sensor. The sensor is very specific to NO because the gas membrane eliminates all ions and other compounds except gases, and the applied electrical potential and electrode material eliminate interferences from other gases such as O2 , CO, and CO2 . The sensor used in this study generates 100-fold higher current than the previous version, providing a more stable signal, with a detection limit of 0.1 nM in comparison with 2 nM of the older sensors (see Note 1) (6, 7). This analytical method is based on the conversion of nitrite to NO in acidic iodide solutions (Fig. 7.1), which can be detected by the sensor. The percent conversion of nitrite to NO in the acidic iodide bath is quantitative as demonstrated by a wide range in linearity between electric current changes and the amount of nitrite added (7).

Fig. 7.1. The conversion of nitrite to nitric oxide in acidic iodide solution (1). For the enzymatic conversion of nitrate to nitrite, nitrate reductase requires the activity of glucose 6-phosphate dehydrogenase (2 and 3).

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Nitrate is not converted directly to NO in the acidic iodide bath. Thus, the conversion of nitrate to nitrite is necessary to measure the total amount of NO formed by the cells. The conversion can be achieved by enzymatic or chemical reductions of which the former is much more efficient. Berkels et al. (6) used nitrate reductase for this purpose (6). In the present method, the reaction of nitrate reductase (Fig. 7.1) is coupled to a glucose 6phosphate dehydrogenase reaction (Fig. 7.1), as in Verdon et al. (8). The conversion of nitrate to nitrite using this coupled enzyme system, termed nitrate reductor (NR), appears to be close to 100% (7). The coupled enzyme system, although its use instead of a single system may look tedious, avoids potential incomplete reduction of nitrate due to NADPH shortage. A big advantage of the current analytical method is that the total NO release from cells in culture can be measured because the degradation products of NO (nitrate/nitrite) accumulate and are stable in the culture media. In the present study, basal NO production is determined in cultured human umbilical vein endothelial cells (HUVECs). In order to examine the specificity and usefulness of the current method, cells are subjected to molecular, biological, and pharmacological treatments leading to the inhibition of nitric oxide synthase 3 (NOS3) expression or enzyme activity, and the resulting changes in NO production are determined.

2. Materials 2.1. Cell Culture

1. HUVECs and endothelial cell growth medium EBM-2 with endothelial growth supplements. 2. Fetal bovine serum and antibiotic antimycotic solution. 3. Cell culture plasticware: 100 mm petri dishes and 6-well plates. 4. 0.2% gelatin. 5. Solutions of trypsin (0.25%) with 0.38 g/L of EDTA.

2.2. Small Interfering RNA (siRNA) Transfection

1. Human NOS3 siRNA (1299001, HSS107237) with nucleotide sequences corresponding to the coding region of a human NOS3 gene transcript (NCBI GenBank accession number, NM_000603.3) and a negative control oligoribonucleotide duplex with scrambled sequences. The nucleotide sequences of the NOS3 siRNA are as follows: 5 -GAA GAG GAA GGA GUC CAG UAA CAC A-3 (sense) and 5 -UGU GUU ACU GGA CUC CUU CCU CUU C-3 (antisense). 2. Lipofectamine RNAiMAX and Opti-MEM (Invitrogen).

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2.3. Assay for NO Production

1. Dulbecco’s modified Eagle’s medium (DMEM) without phenol red formulated to contain 8.3 g/L DMEM powder, 1 g/L D-(+)-glucose, 1 g/L L-glutamine, 110 mg/L sodium pyruvate, and 3.7 g/L sodium bicarbonate (NaHCO3 ). Sterilize by filtration. 2. 100 mM L-NAME) and D-NAME. Sterilize by filtration (see Note 2). 3. Standard solutions of sodium nitrite (NaOH2 ) (0.1– 10 μM). Prepare daily from the stock solution (10 mM). 4. Acidic iodide bath (20 mM potassium iodide (KI) in 0.1 M sulfuric acid (H2 SO4 )). Prepare fresh daily. 5. Nitrate reductor (NR) consists of 0.2 units/mL nitrate reductase, 0.4 units/mL glucose 6-phosphate dehydrogenase, 2 mM glucose 6-phosphate, and 2 μM NADPH in 14 mM Na-P buffer (pH 7.4). Stock solutions of each component are prepared in 14 mM Na-P buffer (pH 7.4) and kept frozen at –80◦ C. Just prior to use, all of these four components are combined in the appropriate volume of Na-P buffer and kept on ice. 6. The NO measuring system: NO model T supplied by Innovative Instruments, Inc. The system comes with software and several types of amperometric NO sensors. The software is installed onto a computer and the NO Model T system is connected to the computer. The AmiNO700 sensor provided has the highest sensitivity for NO detection.

2.4. Western Blotting

1. Cell lysis buffer (10 mM Tris-Cl, 150 mM NaCl, 5 mM EDTA, 0.1% sodium dodecyl sulfate, 1% tritonX-100, 1% deoxycholate pH 7.2) supplemented with 1 mM phenylmethylsulfonyl fluoride and protease inhibitor cocktail. 2. DC protein assay kit (Biorad). 3. Standard solution of bovine serum albumin. 4. Polyvinylidene difluoride membrane. 5. Tween-tris-buffered saline (TTBS) (10×): 2 M NaCl, 200 mM Tris, pH 7.5, 1% Tween-20. 6. Blocking buffer and antibody dilution buffer: 5% (w/v) skim milk in TTBS. 7. Primary antibodies: monoclonal NOS3 antibody, monoclonal β-actin antibody. 8. Secondary antibody: goat antimouse IgG conjugated to horseradish peroxidase. 9. ECL kit.

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3. Methods 3.1. Cell Culture and siRNA Transfection

1. HUVECs are cultured on 0.2% gelatin-coated 100 mm petri dishes at 37◦ C and 5% CO2 in complete growth medium: endothelial cell growth medium (EBM-2) supplemented with endothelial growth supplements, fetal bovine serum (10%), and antibiotics (100 U/mL penicillin, 100 μg/mL streptomycin, and 0.25 μg/mL amphotericin B). 2. Adhering cells are detached using trypsin/EDTA solution and resuspended in growth medium without antibiotics. Cells are seeded on 6-well plates (2 × 105 cells/well) and grown for 1 day in growth medium without antibiotics. 3. Cells at ∼50% confluency are washed with Opti-MEM and treated with 25 nM NOS3 siRNA (or NC) and 2 μL/mL Lipofectamine RNAiMAX in 1 mL Opti-MEM for 3 h. Then, 1.5 mL of growth medium without antibiotics is added and cells are incubated for 1 day. 4. Cells are washed with DMEM, followed by incubation in the same medium for 24 h. Some medium is incubated in wells without cells to provide control media needed for the correction of the endogenous levels of NOx in the culture media (see Note 3). 5. The conditioned media and cells are harvested separately.

3.2. Treatment of Cells with L-NAME and D-NAME

1. Cells are seeded on 6-well plates at a density of 2 × 105 cells/well and grown for 2 days in complete growth medium. 2. Cells are washed with DMEM and treated with 1 mM D-NAME or L-NAME in the same medium followed by a 24-h incubation. Some medium is incubated in wells without cells to provide control media needed for the correction of the endogenous levels of NOx in the culture media. 3. After these treatments, the conditioned media and cells are harvested separately.

3.3. Calibration with the Standard Nitrite Solutions

1. Set up the NO measuring system, in NO model T. 2. The NO sensor is polarized by connecting to the NO system and is immersed in water for a few hours or overnight. 3. The sensor is equilibrated in the 10 mL acidic iodide bath until the background current is stabilized. 4. Add an aliquot (50 μL) of NaNO2 standard solutions to acidic iodide bath and monitor current changes due to NO generation. In the acidic iodide bath, nitrite is quantitatively

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reduced to NO which is detected by the sensor. The peak current increases with the amount of nitrite added. 5. Wait until the current reduces to a background value, and then conduct the second addition. 6. Construct a calibration curve, the current peak vs. the amount of nitrite added. 3.4. Determination of NOx in the Conditioned Media

1. After treatment of the cells, the conditioned medium (including control media) is harvested. 2. An aliquot (100 μL) of the conditioned medium is mixed with a solution (100 μL) of NR to reduce nitrate to nitrite. 3. The mixture is incubated at room temperature for 45 min and kept on ice during analysis. 4. A 50–100 μL sample is added to the acidic iodide bath (10 mL) while stirring with a small magnetic bar. 5. Record the current change generated by an NO sensor (Fig. 7.2).

3.5. Western Blotting

1. After treatments, cells are washed in ice-cold PBS and harvested in 100 μL/well cell lysis buffer. Cell suspensions are incubated on ice for 45 min and centrifuged at 13,400 rcf for 5 min to obtain clear cell lysates. 2. Protein content of cell lysates is determined using a BioRad DC assay kit. 3. Dilute the portion of protein sample to be analyzed 4:1 (v/v) with 5× laemmli buffer and heat for 5 min at 95◦ C in a sealed microcentrifuge tube. 4. Prepare SDS-PAGE gels in an electrophoresis apparatus. 5. Load equal amount of the protein samples (20 μg protein) into wells. Load control wells with molecular weight standards. 6. Connect the power supply and run at 80 V for 40 min and at 120 V for 2 h until the bromophenol blue tracking dye reaches the bottom of the separating gel. 7. When electrophoresis is complete, gels are removed and subjected to electrophoretic protein transfer to a polyvinylidene difluoride membrane at 100 V for 1 h or 30 V for 14 h at 4◦ C. 8. After protein transfer, the membrane is washed in TTBS and then incubated in a blocking solution. 9. The membrane is incubated with a primary antibody (1:1,000) for 2 h with agitation at room temperature, followed by three washes with TTBS.

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500 pA

Blank Medium

siRNA NC

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Cond. media

D-NAME

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No production (nmol.proteining–1.24h–1)

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Fig. 7.2. The panel represents a typical recording demonstrating the molecular and pharmacological inhibition of basal NO production. HUVECs were transfected with a scrambled sequence (NC) control oligonucleotide or with NOS3 siRNA for a 3-h period followed by a 24-h incubation in complete growth medium. Pharmacological inhibition of NOS3 was achieved following a 24-h exposure to 0.1 nM D-NAME or L-NAME. Controls are blank medium to correct for basal levels of NOx and conditioned medium removed from the cells. Quantification of NO production derived from the NO model T recording. Data represent mean ± SEM (n = 3). b’s represent significant differences compared with the corresponding a’s.

10. The membrane is incubated with a secondary antibody (1:3,000) for 1 h with agitation at room temperature, followed by three washes with TTBS. 11. Soak the membrane for 30 s in ECL substrate solution. 12. Remove the membrane, drain, and place face down on a sheet of plastic wrap. 13. In a dark room, place an X-ray film onto the membrane. Expose film for a few minutes. 14. Band intensities are analyzed using the NIH ImageJ program (Fig. 7.3).

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Fig. 7.3. The cell lysates were analyzed by Western blot using antibodies specific to NOS3 or β-actin to show relative expression levels of NOS3 proteins. Blots shown are representatives of at least three independent experiments.

3.6. Statistical Analysis

Statistical analysis was performed by Student’s t-test. A p-value of less than 0.05 based on at least three or more independent experiments was considered to be statistically significant.

4. Notes 1. The apparent sensitivity of the NO sensor used in this study was 1 pA = 15 pM. The detection limit of the sensor was about 0.1 nM which generated 7 pA of current over the noise level (± 2 pA). 2. L-NAME is a specific inhibitor of nitric oxide synthase while its stereoisomer, D-NAME, is not. 3. Due to the sensitivity of the current analytical method, the background NOx level in culture media is easily detected. To correct this value, no cell control media was prepared by incubating media only without cells.

Acknowledgements This work was supported by funding from a National Institute of Health grant HL87012, HL75209, and a World Class University Project (HJ) from the Ministry of Science, Technology and Education of S. Korea. It was also supported by Basic Science Research Program through the National Research Foundation funded by the Ministry of Education, Science and Technology, Republic of Korea (2009-0071171) (YB).

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References 1. Ignarro, L. J., Fukuto, J. M., Griscavage, J. M., Rogers, N. E., Byrns, R. E. (1993) Oxidation of nitric oxide in aqueous solution to nitrite but not nitrate: comparison with enzymatically formed nitric oxide from L-arginine. Proc Natl Acad Sci USA 90, 8103–8107. 2. Patel, R. P., McAndrew, J., Sellak, H., White, C. R., Jo, H., Freeman, B. A., Darley-Usmar, V. M. (1999) Biological aspects of reactive nitrogen species. Biochim Biophys Acta 1411, 385–400. 3. Tsikas, D. (2005) Methods of quantitative analysis of the nitric oxide metabolites nitrite and nitrate in human biological fluids. Free Radic Res 39, 797–815. 4. Kleinhenz, D. J., Fan, X., Rubin, J., Hart, C. M. (2003) Detection of endothelial nitric oxide release with the 2,3-diaminonapthalene assay. Free Radic Biol Med 34, 856–861.

5. Boo, Y. C., Kim, H. J., Song, H., Fulton, D., Sessa, W., Jo, H. (2006) Coordinated regulation of endothelial nitric oxide synthase activity by phosphorylation and subcellular localization. Free Radic Biol Med 41, 144–153. 6. Berkels, R., Purol-Schnabel, S., Roesen, R. (2001) A new method to measure nitrate/nitrite with a NO-sensitive electrode. J Appl Physiol 90, 317–320. 7. Boo, Y. C., Tressel, S. L., Jo, H. (2007) An improved method to measure nitrate/nitrite with an NO-selective electrochemical sensor. Nitric Oxide 16, 306–312. 8. Verdon, C. P., Burton, B. A., Prior, R. L. (1995) Sample pretreatment with nitrate reductase and glucose-6-phosphate dehydrogenase quantitatively reduces nitrate while avoiding interference by NADP+ when the Griess reaction is used to assay for nitrite. Anal Biochem 224, 502–508.

Chapter 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 Abstract Automatic flow-based systems have been applied to assay scavenging capacity against reactive oxygen and nitrogen species, providing analytical tools which can cope with different types and large number of samples. In the present chapter, a flow injection analysis procedure is described for the assessment of peroxynitrite scavenging. A sequential injection analysis procedure is also described for determining the scavenging capacity against the nitric oxide radical. For both systems, reaction between putative antioxidants and the reactive species of nitrogen takes place inside the flow conduits before addition of luminol and further detection of remaining reactive species by chemiluminescence. Key words: Flow injection analysis, sequential injection analysis, scavenging capacity, reactive species of nitrogen, nitric oxide, peroxynitrite, chemiluminescence, automation.

1. Introduction Flow injection analysis (FIA) (1) is an automation tool for chemical analysis that allows the performance of assays which are not feasible when carried out manually, by taking advantage of the reproducible timing attained in these systems. The implementation of methods based on transient light formation, generated by bio- and chemiluminescence (2) is only one example that illustrates the advantages of FIA features. During the 1990s the increased availability of computers in the chemical lab fostered the development of sequential injection analysis (SIA) (3). Based on the same principles of FIA, this new H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_8, © Springer Science+Business Media, LLC 2011

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generation of computer-controlled flow systems offers more flexibility concerning the method development and operation (4). Any changes (sample volume, reagent selection, sample dilution, and reagent-to-analyte ratio) are accomplished via flow programming rather than by physical reconfiguration of the flow path. Therefore, the assessment of antioxidant (AO) capacity using these flow-based systems is rather convenient (5). Hence, the evaluation of scavenging capacity against biologically relevant species, such as reactive species of oxygen (ROS) and reactive species of nitrogen (RNS, including nitric oxide and peroxynitrite), has been proposed (6) with a significant improvement in experimental protocol towards mimicking reaction conditions found in vivo (7). In this context, a versatile and simple FIA system based on chemiluminescence (CL) detection was developed by Sariahmetoglu et al. (8) for the determination of scavenging capacity against several reactive species, including peroxynitrite (ONOO– ) (8). A three-channel manifold (Fig. 8.1) was assembled to accommodate the chemiluminogenic reagent (luminol), the reactive species (ONOO– in this case), and the carrier stream, which transported the injected sample with putative antioxidant properties. Using this manifold configuration, the ONOO– produced offline through the reaction between sodium nitrite and hydrogen peroxide (9) was first mixed with the injected sample. After a residence time of about 1.4 s in the reaction coil, this mixture was subsequently merged with luminol, just before entering in the CL-flow cell. Thus, the FIA design exploits the consumption of ONOO– by antioxidants, which results in the appearance of negative CL signal proportional to the scavenging ability of the compounds (Fig. 8.2). This manifold was also applied to

Fig. 8.1. Schematic representation of FIA system for the chemiluminescence determination of scavenging capacity against peroxynitrite. AO, antioxidant sample; C, MilliQ H2 O as carrier; ONOO– , peroxynitrite standard solution (10–5 M); luminol solution (10–4 M) prepared in carbonate buffer 0.1 M at pH 10; IV, injection valve; P, peristaltic pump; RC, reaction coil (length = 60 cm; volume = 118 μL); CLD, chemiluminescence detector equipped with flow-through cell; PMT, photomultiplier tube; W, waste.

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Fig. 8.2. Schematic representation of (a) inhibition signal profile for increasing concentrations (S1 –S4 ) of test sample with (b) detailed indication of CL signal parameters for calculation of IC50 value in the FIA method for assessing the scavenging capacity against ONOO– .

determination against other reactive species (such as superoxide radical ion, hydrogen peroxide, hypochlorite ion, and hydroxyl radical) by changing the solution fed to one of the flow channels. There are also FIA systems for in vivo nitric oxide (NO) detection (10). These include measurements of physiological NO concentrations present in blood or brain tissues (11) and for pathological NO following acute myocardial infarction (12). A flow-based manifold comprising of a microdialysis probe with CL detection of NO has been described for studying the release of this reactive species in the rat brain following traumatic injury (13). However, none of these systems was applied to the scavenging properties of the compound. Recently, Miyamoto et al. (14) proposed a SIA system for CL determination of NO (and also of O− 2 ) that accommodates both reactions (14). In this system, sample and NOR1 ((±)-(E)-4-methyl-2-[(E)hydroxyimino]-5nitro-6-methoxy-3-hexenamide), a NO donor, were sequentially aspirated into the holding coil (Fig. 8.3). After flow reversal, the two overlapped zones were directed towards the detector while NO was scavenged by antioxidants present in the sample. Subsequently, the remaining NO was measured after addition of luminol from a T connector just before the detector (Fig. 8.4). The system was applied to compounds/enzymes with known activity against this radical, and also to commercial vitamin supplements.

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Fig. 8.3. Schematic representation of SIA system for the chemiluminescence determination of scavenging capacity against nitric oxide radical. AO, antioxidant sample; C, HEPES-NaOH buffer 0.1 M at pH 8.2 as carrier; NOR1, NO donor agent (0.010 μM) prepared in a mixture of DMSO and 0.1 M HCl (50:50, v/v); luminol solution (0.2 mM) prepared in water containing 1% of N,N-dimethylformamide; SV, selection valve; P, peristaltic pump; AP, auxiliary peristaltic pump; HC, holding coil (length = 100 cm; volume = 197 μL); PMT, photomultiplier tube; W, waste.

Fig. 8.4. Schematic representation of analytical signal profile for increasing concentrations (S1 –S3 ) of test sample in the SIA system for the determination of scavenging capacity against NO.

2. Materials 2.1. Automatic Determination of Scavenging Capacity Against Peroxynitrite 2.1.1. Flow Injection Apparatus

Flow injection systems should contain the following components (see Note 1): 1. Multichannel peristaltic pump, with at least three channels, equipped with propulsion tubes (see Note 2).

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2. Rotary injection valve (6-port) equipped with 20 μL loop. 3. Polytetrafluoroethylene (PTFE) tubing (see Note 3). 4. Y-shaped low-pressure connectors and fittings (see Note 4). 5. Chemiluminescence detector equipped with flow-through cell. 6. Chart strip recorder or signal processing unit (see Note 5). 2.1.2. Synthesis of Peroxynitrite Stock Solution

1. Two 10 mL glass syringes. 2. Polytetrafluoroethylene (PTFE) tubing (see Note 3). 3. Y-shaped low-pressure connectors and fittings (see Note 4). 4. Magnetic stirrer. 5. 100 mL glass beaker immersed in ice bath. 6. UV/vis spectrophotometer. 7. 0.6 M sodium nitrite (NaNO2 ) solution: dissolve 1.035 g of NaNO2 in 25 mL of MilliQ H2 O. 8. 0.6 M hydrogen peroxide (H2 O2 ) solution: add 1.5 mL of hydrochloric acid (HCl) (37% m/m, density 1.2 g/mL) to 10 mL of MilliQ H2 O (giving 0.7 M HCl), followed by addition of 1.5 mL of H2 O2 (30% m/m, density 1.1 g/mL), and H2 O to give a final volume of 25 mL. 9. Dissolve 1.2 g of sodium hydroxide (NaOH) in 25 mL of MilliQ H2 O, producing 1.2 M NaOH.

2.1.3. Solutions

1. 0.1 M carbonate buffer (pH 10): dissolve 4.2 g of sodium hydrogen carbonate (NaHCO3 ) in MilliQ H2 O (see Note 6), adding enough 2 M NaOH to give the required pH and making up the volume to 500 mL. 2. 2 M NaOH solution: dissolve 4 g of NaOH in 50 mL of water. 3. 10–3 M luminol stock solution: dissolve 17.7 mg of luminol in 100 mL of 0.1 M carbonate buffer, pH 10 (see Note 7). 4. 10–4 M luminol working solution (see Note 8): tenfold dilution of the 10–3 M luminol stock solution using 0.1 M carbonate buffer. 5. 10–5 M ONOO– solution: appropriate dilution of stock solution in MilliQ H2 O. 6. 10–2 M mannitol solution: dissolve 91.1 mg of mannitol in 50 mL. 7. 10–2 M ascorbic acid stock solution: dissolve 0.1762 g of ascorbic acid in 100 mL of MilliQ water. 8. 10, 50, 100, and 500 μM ascorbic acid solutions.

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2.2. Automatic Determination of Scavenging Capacity Against NO 2.2.1. Sequential Injection Apparatus

Sequential injection systems should contain the following components (see Note 1): 1. Single-channel peristaltic pump equipped with propulsion tubing (see Note 9). 2. Auxiliary single-channel pump. 3. Selection valve (8-port). 4. Polytetrafluoroethylene (PTFE) tubing (see Note 3). 5. Y-shaped low-pressure connectors and fittings (see Note 4). 6. Chemiluminescence detector equipped with flow-through cell. 7. Chart strip recorder or signal processing unit (see Note 5). 8. Hardware and software to operate the selection valve and the main pump (see Note 10).

2.2.2. Solutions

1. 0.1 M HEPES buffer (pH 8.2): dissolve 11.915 g of N-2-hydroxyethyl piperazine-N -2-ethanesulfonic acid in MilliQ H2 O (see Note 6), adding enough 2 M NaOH to give the required pH and making up the volume to 500 mL. 2. 10–4 M NOR1 ((±)-(E)-4-methyl-2-[(E)hydroxyimino]5-nitro-6-methoxy-3-hexenamide) stock solution: dissolve 1.2 mg of NOR1 in 50 mL of a mixture of DMSO and 10–4 M HCl (50:50, v/v). 3. 10–6 M NOR1 intermediate solution: 100-fold dilution of 10–4 M NOR1 stock solution using a mixture of DMSO and 10–4 M HCl (50:50, v/v) as diluent. 4. 10–8 M NOR1 working solution: 100-fold dilution of the 10–6 M NOR1 intermediate solution using a mixture of DMSO and 10–4 M HCl (50:50, v/v) as diluent. 5. 2 × 10–2 M HCl: dilute 1.64 mL of commercial HCl solution (37% m/m, density 1.2 g/mL) with H2 O up to 1 L. 6. 10–4 M HCl: dilute 2 × 10–2 M HCl solution 200-fold with H2 O. 7. 2 × 10–2 M luminol stock solution: dissolve 0.177 g of luminol in 50 mL of N,N-dimethylformamide (DMF) (see Note 7). 8. 2 × 10–4 M luminol working solution (see Note 8): 100fold dilution of the 2 × 10–2 M luminol stock solution using MilliQ H2 O.

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9. 1.0 × 10–2 M ascorbic acid stock solution: dissolve 0.1762 g of ascorbic acid in 100 mL of MilliQ H2 O. 10. 20, 50, 100, 200, and 300 μM ascorbic acid solutions.

3. Methods 3.1. Automatic Determination of Scavenging Capacity Against Peroxynitrite

In the flow injection system, the scavenging reaction against ONOO– takes place at reaction coil RC (Fig. 8.1), where the sample introduced in the system by a rotary valve is mixed with ONOO– solution. After a mean residence time of about 1.4 s at RC (calculated from the flow rate and volume contained in RC), the mixture sample/ONOO– is merged with luminol and directed towards the detector, where a CL signal corresponding to the reaction of the remaining ONOO– with luminol is attained. The total analysis time is less than 20 s, providing a determination throughput of at least 180 per hour.

3.1.1. Synthesis of Peroxynitrite

Stock sodium ONOO– solution is prepared from NaNO2 and H2 O2 by a quenched flow synthesis described by Beckman et al. (9). The original procedure involves a flow network that is simplified. 1. Assemble the system as depicted in Fig. 8.5 (see Note 11). 2. Fill the syringes and the beaker with the respective solutions and immerse all parts in ice. 3. Wait 10 min for temperature stabilization and then press both syringe plungers at the same time as fast as possible (see Note 12), driving NaNO2 and H2 O2 through the synthesis coil into the NaOH solution, where the magnetic stirrer

Fig. 8.5. Schematic representation of experimental flow set-up for the synthesis of peroxynitrite. A, 0.6 M NaNO2 ; B, 0.6 M H2 O2 prepared in 0.7 M HCl; C, 1.2 M NaOH; SC, synthesis coil (volume = 250 μL); D, ice bath; E, magnetic stirrer.

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should be activated. The content of the beaker will turn to yellow and the concentration of ONOO– should be in the range of 170–185 mM (see Note 13). 4. The concentration of ONOO– in the stock solution is assayed by measuring its absorbance at 302 nm and by considering an extinction coefficient of 1,670 M−1 cm−1 (15) (see Note 14). 3.1.2. Operation of Flow Injection System

1. Assemble the flow system as depicted in Fig. 8.1 (see Note 15). 2. Introduce the pumping tubes into the solutions and start the pump (see Note 16). 3. Fill all the lines with the appropriate solutions (see Note 17) and check the flow rate in each channel (see Note 18). Flow rates should be 1.25 mL/min in the channels filled with H2 O and ONOO– solution and 2.5 mL/min in the luminol channel (see Note 19). Wait until a stable baseline signal is attained before starting analysis. 4. Perform the analytical procedure (Section 3.1.3) for all samples. 5. After finishing the analytical procedure, wash the system for 15 min, using H2 O in all channels. Leave all tubing filled with H2 O if the system is to be used within a week; otherwise empty the tubing and disassemble the system.

3.1.3. Analytical Procedure

1. Fill the loop of the rotary valve (load position) with the solution to be tested (see Note 20). 2. Inject the test solution by changing the injection valve position. Wait until a minimum value of CL is attained before returning the injection valve to the load position. 3. Refill the loop and inject the same test solution once again when CL signal returns to its baseline value. Repeat this procedure until achieving 4–5 replicate measurements for the same test solution. 4. To check if the system is working, use ascorbic acid and mannitol as positive and negative controls, respectively (see Note 21). Under these reaction conditions, the IC50 (see Section 3.3) value for ascorbic acid is about 63 μM.

3.2. Automatic Determination of Scavenging Capacity Against Nitric Oxide

For the sequential injection system described here, the scavenging reaction against NO takes place at the holding coil HC (Fig. 8.3) as the sample and NOR1 are sequentially aspirated into the tubing. NOR1 solution is prepared in acidic media and it starts to release NO when mixed with a carrier buffer at pH 8.2. Antiox-

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idant species present in the sample react with NO formed in the HC. The remaining NO is detected by the reaction with luminol after propelling the content of the HC towards the chemiluminescence detector. The total analysis time is around 60 s, providing a determination throughput of 60 per hour. 3.2.1. Operation of Sequential Injection System

1. Assemble the sequential injection system as depicted in Fig. 8.3 (see Note 15). 2. Introduce the pumping tube into the carrier solution, select the waste port in the selection valve, and start the pump (see Note 16) at 1.5 mL/min. Fill the HC completely. 3. Fill the port with NOR1 solution by aspirating a small amount (e.g. 100 μL) of this solution into the HC. 4. Select the waste port and wash the HC (e.g., 400 μL), discarding excess NOR1 solution. 5. Execute the last two steps for antioxidant/test sample, selecting the appropriate port. 6. Select the detector port in the selection valve and fill the tubing connecting the selection valve and the chemiluminescence detector with buffer (see Note 22). During this step, the auxiliary pump must also be activated at 1.0 mL/min. Wait until a stable baseline signal is attained before starting analysis. These operations can be carried out manually or through computer control, depending on the equipment applied. 7. Perform the analytical procedure (Section 3.2.2) for all samples. 8. After finishing the analytical procedure, wash the system using H2 O as a carrier and replace the luminol solution with water. Wash all ports of the selection valve that were used. Leave all the tubing filled with water if the system is to be used within a week; otherwise empty the tubing and disassemble the system.

3.2.2. Analytical Procedure

1. Start the luminol auxiliary pump at 1.0 mL/min. 2. Aspirate 5 μL of the test sample into the HC by selecting the sample port in the selection valve followed by activation of the pump at 0.3 mL/min. 3. Repeat the same operation for the NOR1 solution, selecting the appropriate port. 4. Select the detector port in the selection valve and, reversing the flow at the main pump, propel the HC content towards the detector at 1.5 mL/min for 24 s. Repeat this procedure to obtain 4–5 replicate measurements for the same test solu-

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tion (see Note 23). To change the test solution, follow the procedure indicated in Section 3.2.1. 5. Use ascorbic acid as a positive control (see Note 21). Under these reaction conditions, the IC50 value (see Section 3.3) for ascorbic acid is about 54 μM. 3.3. Sample Analysis

1. The flow systems described here can cope with liquid samples. Hence, any liquid sample can be directly introduced into both systems. Extraction procedures should be performed for solid samples, including a filtration step to remove any particle that would block the flow tubing. 2. Tolerance to organic solvents should be tested by injecting the appropriate dilution of the applied solvent into the flow systems. Signal quenching/enhancement should not be observed (see Note 24). 3. Blank measurements should always be taken to assess: (i) direct reaction between sample components and CL reagents. (ii) formation of luminescent products upon reaction of sample components and reactive nitrogen species (RNS). (iii) intrinsic luminescence from sample components. These measurements are performed by assessing the analytical signal for the highest concentration of test sample, replacing some reagent solutions by their solvents. Hence, in the first case, replace the ONOO– solution with MilliQ H2 O (scavenging capacity against ONOO– ) or NOR1 solution by DMSO and 10–4 M HCl solution (50:50, v/v) (scavenging capacity against NO). In the second case, replace luminol solution with 0.1 M carbonate buffer, pH 10 (scavenging capacity against ONOO– ) or with 1% (v/v) DHF solution (scavenging capacity against NO• ). In the third case, replace both RNS and luminol solutions as described before. For all cases, none or negligible luminescence should be attained in order to exclude interferences on CL signal from sample components. 4. Results are generally expressed as IC50 values defined as the concentration of test sample that provides a 50% inhibitory effect, in this case the 50% decrease of the CL analytical signal attained in the absence of test sample (Figs. 8.2 and 8.4). For the determination of the scavenging capacity against ONOO– , the percentage of inhibition is calculated as:   CLbaseline − CLsample × 100% %inhibition = CLbaseline   negative peak height × 100% = baseline height

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where CLsample is the minimum CL value attained. For the determination of scavenging capacity against NO, as the reactive species is not fed continuously into the detector, the percentage of inhibition is calculated as:  %inhibition =  =

CLblank − CLsample CLblank

 × 100%

blank peak height − sample peak height blank peak height × 100%



where CLblank is the maximum CL value attained for the test sample solvent and CLsample is the maximum CL value attained for the test sample. 5. If liquid sample/extract is introduced undiluted and the CL signal is completely quenched, appropriate dilutions should be taken until partial inhibition of CL signal is attained. Perform at least four different dilution levels to check for linearity in the zone around 50% inhibition (between 20 and 80% inhibition, for instance). If 50% inhibition is not attained, indicate the maximum percentage attained and the respective concentration level (for instance, 35% inhibition for a concentration of 0.2 μg/mL for a given plant extract).

4. Notes 1. Commercial flow injection and sequential injection systems are available from several manufacturers, namely FIAlab Instruments (Bellevue, Washington, USA), Burkard Scientific (Middlesex, UK), Lachat Instruments (Loveland, Colorado, USA), and GlobalFIA (Fox Island, Washington, USA). 2. The flow rate is defined by the rotation speed of the peristaltic pump and by the internal diameter of the pumping tubing. For this particular application, two tubing diameters are necessary in order to provide a given flow rate. This can be attained using Gilson (Villiers-Le-Bel, France) PVC pumping tubes with colour codes black/black (0.76 mm i.d.) and white/white (1.02 mm i.d.). 3. FIA systems usually employ tubing with an internal diameter of 0.8 mm and an external diameter of 1.6 mm. One metre of this tubing contains approximately 502 μL. Tub-

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ing with an internal diameter of 0.5 mm or 1.0 mm can also be applied, but its length should be adapted, considering that 1 m contains 196 and 785 μL, respectively. 4. These connectors and fittings are also commercially available from Omnifit (Cambridge, UK) and Gilson (VilliersLe-Bel, France). 5. FIA systems do not require computer control, with analytical data recorded in strip recorders, where peak height is manually measured. However, nowadays most detection systems have already incorporated data processing features. 6. H2 O used in the preparation of the solutions and buffers should be of high quality, such as H2 O obtained from MilliQ systems (resistivity > 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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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 mbar and is focused onto a liquid-N2 -cooled InSb photodetector. Again, there is a modulation coil around the cell in order to use the Zeeman effect to tune the absorption frequency. The sinusoidal modulation of the absorption and its detection by a lock-in amplifier result in a signal shape similar to the first derivative of the absorption line, showing a zero-crossing at the line center. This error signal is fed back to the QC laser current for frequency stabilization. The detection limit, i.e., the noise-equivalent sensitivity of the spectrometer, was determined by monitoring different dilutions of a certified gas mixture in air, typically NO in nitrogen. The linearity of signal and NO concentration has been checked over by one order of magnitude and the noise-equivalent NO concentration was 25 ppbv (in air), using an averaging time of 300 ms. This time resolution is required, for example, for medical breath analysis. When the averaging time is increased to 10 s, the detection limit achieved is 4 ppbv NO (in air), suitable for monitoring the NO release from liquids. Faraday modulation spectroscopy has also been employed for the simultaneous detection of 14 NO and 15 NO by using a cw distributed feedback QCL operating near 5.4 μm. The simultaneous detection was performed using a fast laser frequency switching between the two isotopologues with a time resolution of 2 s. The isotope ratio has been determined with a precision (1σ ) of 0.052% at 800 s averaging time for 100 ppmv NO gas (44).

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Very recently, a transportable prototype Faraday rotation spectroscopic system based on a tunable external cavity QCL has been developed. A broadly tunable laser source allows targeting the optimum Q3/2 (3/2) molecular transition at 1,875.81/cm of the NO fundamental band. For an active optical path of 44 cm and 1 s lock-in time constant, a minimum NO detection limit (1σ ) of 0.38 ppbv is obtained using a liquid-N2 -cooled InSb photodetector (45). 3.5. Comparison Between Different Methods

In Table 10.1, a detailed comparison between different spectroscopic techniques in terms of advantages and drawbacks is reported. Direct laser absorption techniques give the lowest detection limit (pptv) thanks to long effective optical path lengths and absolute quantitative assessments of species. However, this sensitivity can only be reached with complex and bulky apparatus such as multipass cell and cryogenically cooled low-noise detectors. To obtain longer optical paths (up to several kilometers) high-finesse optical cavity (such as for CRDS and ICOS) can be employed. However, cavity-enhanced approaches present disadvantages for practical applications mostly in terms of setup compactness, robustness, and sensitivity to optical alignment, and thus do not represent the best solution for in situ and online NO detections. An alternative solution may be photoacoustic spectroscopy. This method has the advantages of low sensitivity to optical alignment, ease of use, portability, and high dynamic range. Also, methods based on the Faraday effect have demonstrated good sensitivity; however, they can be employed only for paramagnetic molecules and thus not in multidetection systems. Finally, the best results obtained for the reported spectroscopic techniques are summarized in Table 10.2.

4. Notes 1. QCLs are designed by means of band-structure engineering and grown by molecular beam epitaxy techniques (13). The benefit of this approach is a widely variable transition energy primarily regulated by the thicknesses of the quantum well and barrier layers of the active region rather than the band gap as in diode lasers. 2. Light of known intensity is directed through a gas sample cell and the amount of light transmitted through the sample cell is measured by a detector. If we assume incident

LAS

ppbv or sub-ppbv

Sub-ppbv detection limit thanks to long effective optical path lengths; absolute quantitative assessments of species

Complex apparatus; bulky multipass cells

Techniques

Detection limit

Advantages

Drawbacks Liquid-nitrogencooled detector or lasers; sensitivity of the apparatus to optical alignment

Extremely long effective path length with small cells; intrinsic insensitivity to light source intensity fluctuations

ppbv

CRDS

Liquid-nitrogencooled detector or lasers

Extremely long effective path length with small cells; less critical alignment of the exciting laser beam than CRDS

ppbv

OA-ICOS

Table 10.1 Advantages and drawbacks of QCL-based spectroscopic techniques

Sensitivity to external acoustic noise

Low sensitivity to optical alignment; ease of use and high dynamic range; simple optical alignment

Tens of ppbv

PAS

Can be employed only for paramagnetic molecules

No need for multipass- or high-finesse cavities for detection limit enhancement; no interference from other constituents in the air sample

ppbv

FMS

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Table 10.2 Best results achieved with QCL-based sensors References

Method

λ (μm)

QCL

Detector cooling

Detection limit (ppbv)

Recording time (s)

28

QCLAS

5.3

cw near RT

TE

0.03

30

32

CRDS

5.2

cw cryogenic

Cryogenic

0.7

8

35

OA-ICOS

5.2

cw cryogenic

Cryogenic

0.4

1

40

PAS

5.3

Pulsed RT

–

150

10

42

QEPAS

5.3

EC, cw TE

–

15

5

45

FMS

5.3

cw TE

Cryogenic

0.38

1

light intensity, I0 (x = 0, λ), and transmitted light intensity, I(x, λ), the Lambert–Beer’s law relates the transmitted light to the incident light and the absorption coefficient of the sample, α(λ), as I(x, λ) = I0 ·exp[−α(λ)x], where λ is the wavelength and x is the path length. Concentration is determined from the absorption coefficient. 3. The usefulness of laser spectroscopy in the MIR region is limited by the properties of available IR laser sources, i.e., lead–salt diode lasers, coherent sources based on difference frequency generation, optical parametric oscillators, tunable solid-state lasers, and quantum and interband cascade lasers. In most cases these sources work only at cryogenic temperature or in pulsed mode. 4. The White cell (23, 24) is the oldest arrangement. It consists of two semicircular mirrors, called the “D” mirrors, closely spaced along a common diameter facing a third mirror with two notches in a nearly confocal arrangement. The probe beam enters through one notch and emerges through the other. The number of passes is varied by changing the “D” mirror angle. 5. The Herriott cell (25) has two identical spherical mirrors separated by almost their diameter of curvature (nearly concentric) facing each other. A probe beam launched through a hole in one of the mirrors at an angle to the optical axis completes several number of passes between the mirrors and exits through the same hole (or a hole in the other mirror). The beam bounce pattern and path length are controlled by adjusting the mirror separation. 6. Astigmatic mirror cells (26) are variations of the Herriott cell that spread the light spots over the entire mirror surfaces. This greatly increases the number of spots achievable without overlapping spots and therefore the number

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of passes. This cell type is also more compact and possesses the smallest cell volume to effective path length ratio. In such a multipass cell, the number of passes is typically configured for 90–238, which results in effective optical path lengths from 18 to 210 m for mirror separations of 0.3–0.9 m, thus providing a rising improvement in signal strength. 7. Moeskops et al. (27) used a thermoelectrically cooled cw QCL operating in the wavelength range from 1,854 to 1,874/cm. The collimated radiation was directed into a multipass optical cell (Aerodyne, AMAC-76) with an absorption path length of 76 m and a volume of 300 mL at a pressure of 100 mbars. 8. The NO sensor developed by McManus et al. (28) consists of a cw QCL operating near room temperature at 1,900.07/cm (5.3 μm), a 69-m astigmatic multipass cell, and a TE-cooled IR detector. The pattern of reflections on the astigmatic cell mirrors has been designed to minimize optical interference fringes, which were substantially greater with cw mode than with pulsed operation. 9. Kasyutich et al. (29) integrated in the spectrometer a 1,900/cm cw QCL as source and a thermoelectrically cooled HgCdZnTe photovoltaic detector. The reported results were demonstrated for a 100 averaged spectra acquired within 1.25 s and a single-pass cell of 21 cm base length at about 100 Torr. 10. Cavity-enhanced absorption spectroscopy requires high excitation power (>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 (αL<<1–R), the previous equation becomes:   1−R αL I ≈ 1− I0 2 1−R where

L 1−R α

is the effective path lengths.

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15. The laser source is operated near 5.2 μm with a maximum output power of 80 mW. The 35.5-cm-long high-finesse cavity consists of two dielectric mirrors (R = 99.995% at 5.2 μm, radius of curvature of 6 m) and is characterized by an effective path length of L = 670 m. 16. The PA signal measured by the microphone is given by: S = C · P (λ) · α (λ) where C is the cell constant in the unit of Vcm/W, P the optical power of the laser source, and α the absorption coefficient which is related to the gas concentration (N, number density of molecules) and absorption cross section (σ ) by α = N σ . The cell constant depends on the geometry of the sample cell, the beam profile, the microphone response, and the nature of the acoustic mode. Thus, the sensitivity of a PA sensor can be considerably improved by using resonant photoacoustic sample cells and modulating the laser radiation at a frequency equivalent to an acoustic mode of the cell. As the equation implies, the recorded PA signal can be enhanced also using high-power laser sources emitting in the fundamental absorption region. 17. The laser operated at room temperature in pulsed mode (pulse duration of 42 ns and a duty cycle of 1.4%) with an optical average power of 4 mW at a wavelength around 5.3 μm. The resonant PA cell was a cylindrical stainlesssteel resonator of 120 mm length and 4 mm radius with λ/4 buffer volumes on each side used as acoustic filters. The resonator, designed to be exited in the first longitudinal mode at 1,380 Hz, was equipped with four electret microphones (Knowles EK 3024, 20 mV/Pa), placed at the position of maximum acoustic amplitude to increase the signal-to-noise ratio. 18. The sensor is based on a differential Helmholtz resonant cell formed by two stainless-steel cylindrical volumes (10 cm in length, 5 mm radius) linked by two capillaries (10 cm in length, 2 mm radius) and characterized by a resonance frequency of 315 Hz. The differential measurement, obtained by measuring the PA signal with two microphones (one in each volume), eliminates a great part of the acoustic noise and raises the acoustic signal by a factor of 2. The cell was coupled with a cw DFB-QC laser working at cryogenic temperature in cw mode in the 5.4 μm region. 19. The magnetic field is sinusoidally modulated (with modulation frequency of few kHz). A typical value for the applied alternating magnetic field B is 1 × 10−2 Tesla (RMS),

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which results in a periodic Zeeman shift in the order of ±100 MHz. For demodulation of the signal, a lock-in amplifier is used. Assuming that the polarizers are slightly uncrossed, the output of the lock-in amplifier is proportional to the signal amplitude. The demodulated signal is recorded by a personal computer and the results may be displayed in real time.

Acknowledgements We acknowledge partial financial support from Regione Puglia – Project DM01 related with the Apulian Technological District on Mechatronics-MEDIS. References 1. Nelson, D. D., Shorter, J. H., McManus, J. B., and Zahniser, M. S. (2002) Subpart-per-billion detection of nitric oxide in air using a thermoelectrically cooled midinfrared quantum cascade laser spectrometer. Appl Phys B 75, 343–350. 2. Weber, W. H., Remillard, T. J., Chase, R. E., Richert, J. F., Capasso, F., Gmachl, C., Hutchinson, A. L., Sivco, D. L., Baillargeon, J. N., and Cho, A. Y. (2002) Using a wavelength-modulation quantum cascade laser to measure NO concentration in the parts-per-billion range for vehicle emissions certification. Appl Spectrosc 56, 706–714. 3. Wysocki, G., Kosterev, A. A., and Tittel, F. K. (2005) Spectroscopic trace-gas sensor with rapidly scanned wavelengths of a pulsed quantum cascade laser for in situ NO monitoring of industrial exhaust systems. Appl Phys B 80, 617–625. 4. Kharitonov, S. A. and Barnes, P. J. (2000) Clinical aspects of exhaled nitric oxide. Eur Respir J 16, 781–792. 5. van Herpen, M. M. J. W., Bisson, S. E., Ngai, A. K. Y., and Harren, F. J. M. (2004) Combined wide pump tuning and high power of a continuous-wave, singly resonant optical parametric oscillator. Appl Phys B 78, 281–286. 6. Bisson, S. E., Armstrong, K. M., Kulp, T. J., and Hartings, M. (2001) Broadly tunable, mode-hop-tuned cw optical parametric

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26. McManus, J. B., Kebabian, P. L., and Zahniser, M. S. (1995) Astigmatic mirror multipass absorption cells for long-path length spectroscopy. Appl Opt 34, 3336–3348. 27. Moeskops, B. W. M., Critescu, S. M., and Harren, F. J. M. (2006) Sub-part-per-billion monitoring of nitric oxide by use of wavelength modulation spectroscopy in combination with a thermoelectrically cooled, continuous-wave quantum cascade laser. Opt Lett 31, 823–825. 28. McManus, J. B., Nelson, D. D., Herndon, S. C., Shorter, J. H., Zahniser, M. S., Blaser, S., Hvozdara, L., Muller, A., Giovannini, M., and Faist, J. (2006) Comparison of cw and pulsed operation with a TE-cooled quantum cascade infrared laser for detection of nitric oxide at 1900 cm−1 . Appl Phys B 85, 235– 241. 29. Kasyutich, V. L., Holdsworth, R. J., and Martin, P. A. (2008) Mid-infrared laser absorption spectrometers based upon all-diode laser difference frequency generation and a room temperature quantum cascade laser for the detection of CO, N2 O and NO. Appl Phys B 92, 271–279. 30. Romanini, D., Kachanov, A. A., Sadeghi, N., and Stoeckel, F. (1997) CW cavity ring down spectroscopy. Chem Phys Lett 264, 316–322. 31. Paldus, B. A., Harb, C. C., Spence, T. G., Zare, R. N., Gmachl, C., Capasso, F., Sivco, D. L., Baillargeon, J. N., Hutchinson, A. L., and Cho, A. Y. (2000) Cavity ringdown spectroscopy using midinfrared quantum-cascade lasers. Opt Lett 25, 666–668. 32. Kosterev, A., Malinovsky, A. L., Tittel, F. K., Gmachl, C., Capasso, F., Sivco, D. L., Baillargeon, J. N., Hutchinson, A. L., and Cho, A. Y. (2001) Cavity ringdown spectroscopic detection of nitric oxide with continuouswave quantum-cascade laser. Appl Opt 40, 5522–5529. 33. Menzel, L., Kosterev, A. A., Curl, R. F., Tittel, F. K., Gmachl, C., Capasso, F., Sivco, D. L., Baillargeon, J. N., Hutchinson, A. L., Cho, A. Y. et al. (2001) Spectroscopic detection of biological NO with a quantum cascade laser. Appl Phys B 72, 859–863. 34. Bakhirkin, Y. A., Kosterev, A. A., Roller, C., Curl, R. F., and Tittel, F. K. (2004) Midinfrared quantum cascade laser based off-axis integrated cavity output spectroscopy for biogenic nitric oxide detection. Appl Opt 43, 2257–2265. 35. Silva, M. L., Sonnenfroh, D. M., Rosen, D. I., Allen, M. G., and O’Keefe, A. (2005) Integrated cavity output spectroscopy measurements of NO levels in breath with a

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Chapter 11 Determination of In Vivo Nitric Oxide Levels in Animal Tissues Using a Novel Spin Trapping Technology Anatoly F. Vanin and Alexander A. Timoshin Abstract It has been established that microdialysis ensured by the passage of aqueous solutions of Fe3+ complexes with N-methyl-D-glucamine dithiocarbamate (MGD) through fine dialysis fibers permeable for compounds with molecular weights below 5 kDa. These fibers can be implanted into heart, liver, and kidney tissues, enabling effective binding of Fe3+ –MGD complexes to nitric oxide generated in interstitial fluids of narcotized rats in vivo. Subsequent treatment of dialyzate samples (60 μL) with sodium dithionite favors conversion of newly formed diamagnetic NO–Fe3+ –MGD complexes into electron paramagnetic resonance-detectable NO–Fe2+ –MGD complexes. The basal levels of NO determined from the concentrations of the complexes in the respective tissues are similar (1 μM). The microdialysis data suggest that treatment of rats with a water-soluble analogue of nitroglycerine or a dinitrosyl iron complex with thiosulfate induces a long-lasting (>1 h) increase in the steady-state level of NO in animal tissues. This novel technology can be used for comparative analyses of production rates of NO and reactive oxygen species when using iron–dithiocarbamate complexes and spin traps for reactive oxygen species, respectively. Key words: Cardiac ischemia, electron paramagnetic resonance, microdialysis, nitric oxide, reactive oxygen species, NO spin trapping.

1. Introduction More than 20 years ago, a team of investigators led by one of the authors of the present communication proposed to use bivalent iron complexes with diethyldithiocarbamate (DETC) for spin trapping nitric oxide (NO) produced by animal tissues and cell cultures (1). The binding of NO to these complexes stimulates the production of paramagnetic mononitrosyl iron complexes H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_11, © Springer Science+Business Media, LLC 2011

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(MNIC–DETC) detectable by the EPR method (see Note 1). The intensity of the EPR signal corresponds to the concentration of MNIC and, as a consequence, to the amount of NO incorporated into them. Further experiments showed that enhanced incorporation into MNIC–DETC of NO produced by various biosystems can be related to the high affinity of Fe2+ –DETC complexes for NO molecules (2–4). Iron complexes with DETC, similar to MNIC–DETC, are hydrophobic and localized in hydrophobic compartments of the cell (e.g., cell membranes), but are never secreted into circulating blood or excreted with urine. As a result, MNIC–DETC are retained within the composition of organs and tissues where they are accumulated in large amounts until a steady-state level is reached after 30–60 min, provided that their production and decomposition rates are similar. The main reason for their decomposition is the interaction of NO incorporated into MNIC– DETC with superoxide anions (4, 5) and DETC hydrolysis, which yields carbon disulfide and ethanolamine. The ever-increasing popularity of Fe2+ –DETC complexes as effective spin traps for NO enables determination of ex vivo accumulation of NO in normal and pathologically changed animal organs by the L -arginine-dependent pathway. Furthermore, this also allows the acquisition of valuable information about the fine mechanisms of NO production in animal tissues and cell cultures (2–4, 6–16). The use of EPR spectroscopy in the L-band allows monitoring of MNIC–MGD formation in vivo, particularly in experiments with mice and rats placed into the cavity of a radiospectrophotometer (8, 17, 18). However, the results of these studies are limited, being confined to only general information about NO production in a whole animal body without giving its estimation in the tissues separately. More detailed information about NO generation in various animal tissues in vivo can be derived from EPR imaging studies, but the practical application of EPR tomography is limited because of the complexity and high costs of the respective equipment (19–21). Water-soluble iron–dithiocarbamate complexes, e.g., iron complexes with N-methyl-D-glucamine dithiocarbamate (MGD) able to bind NO, were first obtained in the 1990s (22, 23). In animal models, these complexes which are similar to MNIC circulate in the blood and intercellular fluids but never penetrate the cell (see Note 2). The hydrophilic nature of iron complexes with MGD and other ligands makes them helpful tools in the study of NO production by isolated NO synthases in aqueous solutions, plants, and various cell cultures (24–31). One of the disadvantages of water-soluble iron complexes with dithiocarbamates for determining NO levels in animal tissues is their fast excretion in urine (see Note 3). Therefore, these

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compounds are administered to animals at much higher concentrations than their hydrophobic analogues (29). The distribution of water-soluble MNIC in animal tissues can be estimated under ex vivo conditions, i.e., after sacrifice and extraction of internal organs. Here we describe a novel approach for determining NO levels in various organs in vivo where water-soluble iron complexes with MGD are used as effective spin traps for NO. Binding of NO to these spin traps is achieved through microdialysis in fine dialysis fibers permeable for only small molecules and implanted into various animal tissues (heart, liver, kidney, etc.). In relation to in vivo NO estimation in lung tissue, we pass exhaled air through solutions of iron complexes with MGD that binds to NO. The results of our previous studies (32) and our most recent (unpublished) data provide an illustrative example of the practical application of these novel technologies.

2. Materials 2.1. Iron Complexes with MGD

1. Sodium N-methyl-D -glucamine dithiocarbamate (MGD) synthesized using the previously described procedure (33). 2. Ferrosulfate (Fluka, Switzerland). 3. 150 mM phosphate buffer rN 7.4 (molar ratio 10:1). 4. Sodium dithionite.

2.2. In Vivo Models

1. 8–11-mm-long dialysis fibers (Codis Dow, Belgium) with external and internal diameters 0.25 and 0.1 mm, respectively, and a 5-kDa molecular weight cut-off. 2. Male Wistar rats weighing 300–400 g. 3. Ketamine (100 mg/kg body weight). 4. Catheters. 5. Artificial pulmonary ventilation unit. 5. Liquid nitrogen. 6. NO spin traps 0.5 mL. 7. Three-channel syringe pump (CMA-100, Sweden).

2.3. EPR Assays

1. X-band Varian E-109E radiospectrometer. 2. Thiosulfate. 3. Dimethylsulfoxide. 4. Isosorbide dinitrate Isoket 0.1%.

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3. Methods 3.1. Aqueous Solutions of Iron Complexes with MGD

1. Dissolve MGD and ferrosulfate [Fe(SO4 ) × 7H2 O] in 150 mM PBS at rN 7.4 (molar ratio 10:1) to a final salt concentration of 30 mM. Leave for 1–2 min until the solution turns brown. This is a result of the oxidation of Fe2+ ions within the composition of Fe2+ (MGD)2 complexes to the trivalent (Fe3+ ) state (34–39) (see Note 4). 2. Convert the diamagnetic NO–Fe3+ (MGD)2 complex into the paramagnetic (NO–Fe2+ (MGD)2 ) form, using sodium dithionite (a reducing agent) at a final concentration of 5 mM (35–40).

3.2. In Vivo Studies

1. Anesthetize male Wistar rats with ketamine (100 mg/kg). 2. Insert the catheters into the right carotid artery and the right jugular vein. 3. Perform a tracheotomy to attach an artificial pulmonary ventilation unit. 4. Perform a thoracotomy and a laparotomy for the implantation of the dialysis fibers using a needle into the myocardium, liver, and kidney tissues. 5. Pump solutions of Fe3+ (MGD)2 complex through the fibers using a three-channel syringe pump (Fig. 11.1) (see Note 5). 6. Wash dialysis fibers for 10 min with a standard solution of the Fe3+ (MGD)2 complex at a flow rate of 10 μL/min, followed by a flow rate of 3 μL/min for a further 10 min.

Fig. 11.1. A scheme of myocardial microdialysis. 1, dialysis fiber; 2 and 3, dialyzate inflow and outflow silicone tubes; 4, syringe of the microperfusion pump. (Reproduced from Ref. (32) with permission from Elsevier Science.)

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7. Using microtubes, collect the serial effluents from the dialysis fibers and freeze in liquid nitrogen for EPR assays. Each collection cycle lasts 20 min. 8. Fill a flow-through cell with a standard solution of NO spin traps 0.5 mL and insert into the artificial outlet of the pulmonary ventilation outlet. Bubble exhaled air through the NO spin trap solution for 20 min. Freeze and store the dialyzates in liquid nitrogen and thaw immediately before EPR assays. 3.3. EPR Assays

The intensity of EPR signals of paramagnetic MNIC–MGD complexes formed in dialyzates or spin trap solutions is high enough to enable detection on a X-band Varian E-109E radiospectrometer (USA) at ambient temperature. The EPR signal (g = 2.04) is characterized by a triplet hyperfine structure (HFS) with HF splitting 1.2 mT as a result of HF interaction of the unpaired electron with the nucleus of the nitrogen atom of the nitrosyl ligand. The EPR signals recorded in myocardial tissue dialyzates and Fe–MGD solutions bubbled with exhaled air are shown in Fig. 11.2a,b, respectively. 1. Collect 60 μL of the spin trap solution dialyzates after 20 min passage through the dialysis fibers. 2. Determine the concentration of MNIC–MGD by the double integration of the EPR signal. 3. Generate a reference sample by combining Fe2+ + thiosulfate with gaseous NO (41) at the Fe2+ thiosulfate molar ratio 1:20. This should ensure maximal incorporation of iron into

Fig. 11.2. Panel a: EPR spectra of dialyzate samples from the myocardium. 1 and 2, 2–20 min passage of standard solution of Fe–MGD in phosphate buffer, pH 7.4 (1, without sodium dithionite; 2, after addition of sodium dithionite powder). 3, Passage of a phosphate buffer through dialysis fiber in the absence of Fe–MGD following addition of Fe–MGD and sodium dithionite to the dialyzate sample. Panel b: EPR spectrum of a Fe–MGD solution after 20-min bubbling with exhaled air followed by dithionite addition. All the EPR spectra were recorded at ambient temperature. (Reproduced from Ref. (32) with permission from Elsevier Science.)

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the DNIC producing an EPR signal with a narrow symmetric singlet (g = 2.03) at ambient temperature. The concentration of paramagnetic MNIC–MGD in heart and lung tissues was equal to 1.0 ± 0.1 and 2.0 ± 0.2 μM/L, respectively, being commensurate with the levels of NO formed from endogenous sources (see Notes 6, 7, 8, and 9; Fig. 11.3). Subsequent treatment of experimental animals with the NO synthase inhibitor resulted in the disappearance of paramagnetic MNIC–MGD complexes from the dialyzates.

Fig. 11.3. Panel a: Times-courses of MNIC–MGD concentration in dialyzate samples corresponding to heart, liver, or kidney. Forty minutes after the onset of microdialysis – start of Isoket infusion (means ± SEM from four to five independent measurements). Panel b: Time-courses of MNIC–MGD concentration in the sample of phosphate buffer with Fe–MGD after 20-min bubbling with exhaled air. Forty minutes after the onset of experiment – DNIC with thiosulfate injection or start of Isoket infusion (means ± SEM from four to five independent measurements). Panel c: Times-courses of MNIC–MGD concentration in dialyzate samples corresponding to heart, liver, or kidney. Forty minutes after the onset of microdialysis – injection of DNIC with thiosulfate (means ± SEM from four to five independent measurements). (Reproduced from Ref. (32) with permission from Elsevier Science.)

Similar EPR signals of MNIC–MGD were recorded in liver and kidney tissue samples; their intensity corresponded to their concentration (~1.0 μM/L) (Fig. 11.3). Using the novel spin trapping technology, it was established that the levels of NO generated from endogenous sources in the heart, liver, and kidney tissues of experimental rats were very similar. At the same time, their comparison with the NO content in lung tissue is incorrect because of the difference between the conditions of NO detection by microdialysis and spin trapping in solutions bubbled with exhaled air. Substitution of an aqueous solution of Fe3+ (MGD)2 for a solution of Fe3+ –DETC in an organic solvent (e.g., dimethylsulfoxide) in the course of microdialysis resulted in the penetration of the solvent into interstitial fluids, which might provoke significant disturbances in tissue metabolism. The use of the novel technology for determining in vivo NO levels in animal tissues enabled statistically significant evaluation of changes in NO levels in animals treated with exogenous substances, such as the water-soluble nitroglycerine derivative

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Isoket, DNIC, and RS-NO. Exogenous NO donors were injected into animals 40 min after the beginning of passage of the spin trap solutions through the dialysis fibers or after the beginning of bubbling the solutions with exhaled air. These findings suggest that the treatment of rats with DNIC + thiosulfate increases NO levels in the heart, lung, liver, and kidney tissues. In the case of Isoket infusion, the increase in NO is especially apparent in myocardial tissue, most probably due to its more selective accumulation in the myocardium (see Note 10). Microdialysis allowed comprehensive analysis of in vivo levels of NO and reactive oxygen species (ROS), e.g., under conditions of local ischemia–reperfusion of the myocardium. ROS levels are determined with the help of any currently known ROS-specific spin traps, e.g., 5,5-dimethyl-1-pyrroline-N-oxide (DMPO). The EPR signals recorded in DMPO solutions in phosphate buffer after their passage through the dialysis fibers implanted into ischemic or nonischemic (intact) zones of animal myocardium are shown in Fig. 11.4. In both solutions, EPR signals were characterized by four superfine structure (SFS) components at the amplitude ratio 1:2:2:1 characteristic of paramagnetic adducts of DMPO–OH (Fig. 11.5) (42). DMPO–OH is a result of the generation of hydroxyl radicals formed when the dialysis fibers react with

Fig. 11.4. Panels a and b: Time-course of DMPO–OH concentration in dialyzate samples corresponding to intact or ischemic myocardial zones in control and injected with DNIC with glutathione (DNIC–GS) rats, respectively. Panels c and d: Time-course of MNIC–MGD concentration in dialyzate samples corresponding to intact or ischemic myocardial zones in control and injected with DNIC with glutathione (DNIC–GS) rats, respectively.

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Fig. 11.5. The EPR signal of DMPO–OH. The record was made at ambient temperature.

endogenous hydrogen peroxide (H2 O2 ). However, the DMPO– ON could also be a product of spontaneous conversion of the DMPO–OOH adduct (42). The time-dependent changes in the intensity of EPR signals of DMPO–ON in the ischemic and intact zones of the myocardium are shown in Fig. 11.4a. Tissue ischemia was induced by a 40-min occlusion of the anterior descending artery with a subsequent 60-min reperfusion of the risk zone. The notable increase in the DMPO–ON level in the risk zone might reflect the increase in the ROS content in the interstitial zone of the ischemic myocardium. In the intact zone, the ROS level did not change. Similar changes were established for the MNIC–MGD content in the ischemic and intact zones of the myocardium (Fig. 11.4c). The increase in the NO level in the ischemic zone begins with the onset of occlusion. After a single (bolus) treatment of DNIC with glutathione (i/v, 3.1 μM/kg) at the onset of local myocardial ischemia, DMPO–ON and MNIC– MGD increased in the ischemic zone along with simultaneous slight increase in the MNIC–MGD concentration in the intact zone (Figs. 11.4b,d). The suppression of the DMPO–ON formation in the ischemic zone can be attributed to the antioxidant effect of DNIC with glutathione, which fulfils the function of an ROS scavenger (43, 44). Such a decrease can be beneficial during subsequent reperfusion of the ischemic zone, being consistent with the protective effect of DNIC on myocardial infarction induced by ischemia–reperfusion (45, 46). As regards the increase in MNIC– MGD levels in the intact zone, it might be due to the interaction of Fe–MGD with exogenous DNIC, with glutathione used as NO donors. The decrease of MNIC–MGD in the ischemic zone is more difficult to interpret (Fig. 11.4d). Most probably, NO production in this particular zone was provoked by local

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acidosis, which in the absence of O2 was accompanied by enzymatic and nonenzymatic reduction of NO2 – and S-nitrosothiols and a concomitant release of NO. On entering this zone, exogenous DNIC were decomposed and the iron ions and glutathione were liberated with subsequent accumulation in the myocardial tissues. However, the binding of the iron ions and glutathione to NO yielded DNIC. This process resulted with interactions with proteins blocking penetration of the dialysis fibers subsequently resulting in a decrease in NO and, correspondingly, MNIC–MGD levels. 3.4. Postscriptum

This technology eliminates numerous problems with adequate interpretation of genuine mechanisms underlying the formation of EPR-detectable MNIC with dithiocarbamates, with MGD. The results of our pioneering studies (1, 2) and those reported by other investigators suggest that the formation of paramagnetic MNIC with such ligands is a result of binding of NO to Fe2+ complexes with dithiocarbamates in animal tissues. However, treatment of animals with Fe2+ –MGD complexes demonstrated that in these complexes iron is rapidly oxidized to the trivalent state. MGD does not compete with other intracellular iron-chelating agents, such as citrate. As a result, in the absence of NO, its direct spin traps (iron–MGD complexes) are not detected in animal tissues. The appearance of NO in such biosystems is a prerequisite to the binding of iron. The affinity of iron for MGD sharply increases at the expense of NO binding; the MNIC– MGD formed contain a Fe3+ adduct with NO, which manifests high oxidative capacity and are easily converted into paramagnetic NO–Fe2+ –(MGD)2 complexes detectable by the EPR method. Therefore, the original hypothesis on exogenous Fe2+ –(MGD)2 as direct NO traps is unrealistic. Our studies on the use of iron– dithiocarbamates for spin trapping NO in various biosystems (1, 2) cannot be regarded as a claim of priority for the novel technology, since there was failure to get a deeper insight into fine mechanisms underlying the formation of EPR-detectable MNIC with dithiocarbamates. While applying the microdialysis method, we deliberately introduced preoxidized iron complexes with MGD into dialysis fibers. It is not excluded that part of these complexes degraded as a result of which iron was transferred to endogenous citrate, which then penetrated inside the dialysis fibers. However, this process only weakly affects the formation of MNIC–MGD due to the penetration of NO into the dialysis fibers. Under these conditions, low-molecular agents able to reduce oxidized MNIC– MGD to the paramagnetic state can penetrate inside the dialysis fibers in only insignificant amounts, which brings forth the necessity of additional treatment of diamagnetic MNIC with exogenous reducing agents of which sodium dithionite seems to be the most efficient.

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4. Notes 1. Water-insoluble iron and DETC complexes are formed in animal tissues and cell cultures as a result of the separate sequential addition of aqueous solutions of DETC and Fe2+ citrate. Aqueous solutions of DETC are administered to animals by i.p., while iron complexes are delivered by i.m. injection to the leg. DETC molecules localized in cell membrane bind to exogenous iron to form NO traps, which represent iron complexes with DETC. The bivalent iron in these complexes is rapidly oxidized to a trivalent state by oxygen. As in the case of the reduced forms of the complexes, the binding of NO to Fe3+ DETC is effective and yields EPR-undetectable diamagnetic MNIC–DETC, the greater part of which is reduced by endogenous reducing agents, e.g., thiols, to the paramagnetic EPR-detectable state. Reduction of all diamagnetic MNIC–DETC is especially effective after treatment of isolated tissues and cells with sodium dithionite (35, 38–40). 2. In contrast to the separate introduction of DETC and iron complexes into animal organisms and cell cultures, Fe–MGD are administered to animals in the preferred state (e.g., as aqueous solutions) either i.p. or i.v. (animal studies), or are added directly to the incubation medium (cell cultures) (22–31). In such solutions, bivalent iron forms a complex with MGD and is rapidly oxidized to the trivalent state by atmospheric oxygen; the binding of NO to iron is accompanied by the formation of EPR-undetectable diamagnetic MNIC–MGD complexes. Subsequent reduction of the latter to the paramagnetic EPR-detectable state takes place after the addition of endogenous and exogenous (dithionite) reducing agents (38–40). 3. A comparison of iron complexes with DETC and MGD for spin trapping NO in different mouse organs ex vivo demonstrated that hydrophobic iron complexes with DETC manifest a higher binding capacity (12, 27). When iron complexes with DETC or MGD were added to mice in equimolar (125 μM/kg) amounts, the content of MNIC– DETC formed in the liver during 30 min exceeded that of MNIC–MGD by three-fold (27). Conversely, blood plasma levels of MNIC–MGD were notably higher. Moreover, small amounts of MNIC–MGD were found in the urine where MNIC–DETC were virtually absent. After 60 min, the MNIC–DETC content in animal tissues did not change, while MNIC–MGD disappeared completely. Thirty minutes after treatment of mice with 17 μM/kg of

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exogenous ready-to-use MNIC–MGD, their total level in different tissues was similar, whereas 60 min thereafter they disappeared from the animal organs, being detected only in the urine. The fact that urine content of MNIC–MGD was 50 times lower than that of exogenous MNIC–MGD provides conclusive evidence that the greater part of these complexes were destroyed, most probably, under deleterious effect of superoxide anions. 4. The oxidized Fe3+ (MGD)2 complex (30 mM), which can effectively bind NO (34–39), was used as a standard solution for spin trapping of NO in animal studies where NO levels were determined using both microdialysis and NO detection in exhaled air. 5. The permeability of dialysis fiber walls enabled diffusion of only low-molecular compounds, e.g., endogenous NO molecules, NO2 – , low-molecular S-nitrosothiol (RS-NO), and dinitrosyl iron complexes (DNIC), into the dialysis fibers (4). 6. During passage of Fe3+ (MGD)2 solutions through the dialysis fibers, part of them are released through the porous wall into interstitial tissues, which diminishes the efficiency of NO spin trapping. After treatment of experimental solutions with the exogenous NO donor GS-NO (both before and after passage through dialysis fibers), the concentration of Fe3+ (MGD)2 complexes decreases three-fold. Presumably, MNIC–MGD are released from the fibers in a similar way. As a result, the level of EPR-detectable MNIC–MGD in effluent samples (60 μL) after their passage through the fibers is three times lower than their total concentration in animal tissues. 7. From Note 6. It follows that the concentration of MNIC– MGD in animal tissues determined by the microdialysis method was as low as 30% of the total NO content in interstitial tissue. Seemingly, the remainder was bound to iron complexes with MGD released from the fibers. Determination of NO content in lung tissue by the amount of Fe3+ (MGD)2 formed by bubbling of the solution with exhaled air has proven to be efficient. 8. The MNIC–MGD formation in dialysis fibers is exclusively due to the ability of Fe3+ (MGD)2 to bind free NO molecules. The contribution of other lowmolecular-weight NO donors (NO2 – , endogenous DNIC, S-nitrosothiols) to this process is negligibly small. Evidence for this hypothesis can be derived from the results of our recent studies in which the solution contained only the phosphate buffer, and Fe3+ (MGD)2, where dithionite was

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added to the solution only after its passage through the dialysis fibers (32). Under these conditions, NO2 – , DNIC, and S-nitrosothiols accumulated in the phosphate buffer might favor the formation of EPR-detectable MNIC– MGD. At the same time, the total lack of these complexes in the solution is explicit of their insignificant role in MNIC–MGD formation. 9. On entering dialysis fibers, superoxide anions do not induce any decomposition of MNIC–MGD. In aqueous solutions, the rate of formation of their adducts with O2 is described by the equation: V = k[MNIC-MGD] [O2 – ], where k = 3 × 10–7 /M/s (4). By substituting MNIC– MGD = 1 μM (maximum concentration of the complexes) and O2 - = 10–9 M (steady-state concentration of the superoxide in the tissues) (47) in the equation, we obtain: V = 3 × 10–8 /M/s. Considering that the reaction time did not exceed 0.4 s coupled with the internal volume of the fibers (0.02 μL) and a flow rate of the spin trap of 3 μL/min, the concentration of the MNIC–MGD adducts and O2 – formed during the same time interval was 1.2 × 10–9 M, i.e., less than that of MNIC–MGD by three orders of magnitude. The optimum concentration of MNIC was determined during the first 20–30 min after NO trap injection. During this time interval, the yield of MNIC–MGD adducts with the superoxide (V × 1.2 × 103 s) reached 3.6 × 10–5 M, which might significantly interfere with the tissue level of MNIC–MGD. Therefore, the reduction of the time of MNIC–MGD exposure to superoxide anions in the course of microdialysis can be regarded as the main reason for the failure to establish any effect of O2 – on the level of MNIC–MGD detectable by the EPR method. 10. High steady-state levels of NO in animal tissues after treatment with Isoket and DNIC did not change for at least 80 min. In contrast, the level of protein-bound DNIC in rats treated with DNIC with thiosulfate decreased threefold within the same period (32).

Acknowledgement This work was financially supported by the Russian Foundation for Basic Research (grants No 08-04-00665a and 09-0400886a).

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Determination of In Vivo NO Levels in Animal Tissues 44. Shumaev, K. B., Gubkin, A. A., Serezhenkov, V. A., Lobysheva, I. I., Kosmachevskaya, O. V., Ruuge, E. K., et al. (2007) Interaction of reactive oxygen and nitrogen species with albumin- and methemoglobin-bound dinitrosyl iron complexes. Nitric Oxide Biol Chem 18, 37–46. 45. Pisarenko, O. I., Shulzhenko, V. S., Studneva, I. M., Vanin, A. F., Chazov, E. I. (2008) Biochemical mechanisms of dinitrosyl iron complex action on ischemic rat heart. Izvestiya Ross Acad Nauk ser Biol No 1, 1–5.

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Section II Nitric Oxide Generation

Chapter 12 β Cell Protection by Inhibition of iNOS Through Lentiviral Vector-Based Strategies Sean O. Hynes, Cillian McCabe, and Timothy O’Brien Abstract Cytoprotective gene transfer to pancreatic islet β cells may prove useful in preventing their destruction and prolonging islet graft survival after transplantation in patients with type 1 diabetes mellitus. A host of therapeutically relevant transgenes may potentially be incorporated into an appropriate gene delivery vehicle and used for islet modification. To examine this, we utilised a robust model of cytokine-induced β cell pathophysiology. Using this model, it is clear that antioxidant gene transfer confers no cytoprotective benefit. In contrast, we demonstrated that gene-based approaches to inhibit the activation of NF-κB following cytokine exposure harbours therapeutic utility in preserving islet β cell viability in the face of cytokine toxicity. We identified that NF-κB-dependent induction of iNOS is a critical determinant of β cell fate following cytokine exposure. Having identified the pivotal role of iNOS activation in cytokineinduced β cell pathophysiology, lentiviral vectors may be used to efficiently deliver small interfering RNA molecules to confer efficient iNOS gene silencing. We have shown that lentiviral vector-based shRNA delivery holds significant promise in preserving β cell viability following cytotoxic cytokine exposure. Key words: Diabetes mellitus, pancreatic islet cells, iNOS, small interfering RNAs, NF-κB, gene silencing, gene therapy.

1. Introduction The destruction of pancreatic islet β cells through autoimmune damage is essential to the pathogenesis of type 1 diabetes mellitus (T1DM) (1). The aetiology of this autoimmune response is not clear. Exogenous treatment of T1DM with insulin is at present the most clinically relevant therapy. However, this treatment does not prevent secondary complications such as retinopathy, nephropathy, or peripheral vascular disease (2). Alternative H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_12, © Springer Science+Business Media, LLC 2011

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strategies for maintaining physiological-based control of insulin include pancreatic and islet cell transplantation, although pancreatic transplantation is associated with significant mortality and morbidity (3). Successful islet cell transplantation has already been demonstrated in both preclinical and clinical settings (4, 5). The introduction of the Edmonton protocol (5) for islet cell transplantation demonstrated 80% independence of insulin, 1-year post intervention. However, this is difficult to maintain and drops precipitously to less than 10% after 5 years (6). Transplanted islet cells fail due to both non-specific inflammatory reactions and specific autoimmune reactions. The principal proinflammatory cytokines which contribute to the destruction of β cells includes interleukin-1β (IL-1β), interferonγ (INF- γ), and tumour necrosis factor-α (TNFα) (7). It has been shown that IL-1β can induce β cell destruction in a nitric oxide (NO)-dependent manner (8). IL-1β stimulates inducible nitric oxide synthase (iNOS) from activated macrophages. INOS produces both NO which is specifically cytotoxic to β cells and if uncoupled may produce reactive oxygen species capable of further damaging β cells through independent pathways (9). NF-κB contributes to this pathophysiology through its activation by both IL-1β and TNFα which allow phosphorylation and degradation of inhibitor κB (I- κB). Once activated NF- κB binds to DNA and modulates gene expression rapidly contributing to β cell degradation. A strategy which utilises gene therapy techniques to inhibit iNOS would be valuable in enhancing the surviving fraction of transplanted islet cells. Also, since islet cells can survive ex vivo it would be possible to genetically manipulate these cells prior to transplantation. Gene therapy techniques can achieve this through the use of short hairpin interfering RNAs (shRNAs) to silence iNOS expression. Initial work showed that delivery of shRNAs was capable of silencing expression of the tumour suppressor gene p53 in mice, allowing tumour formation (10). Previous studies have shown the promise of RNA interference in modulating islet cell expression profiles (11–16). Delivery of shRNAs to islet cells may be achieved through gene therapy techniques. In particular, the use of adenoviral and lentiviral vectors has shown to be effective in transducing islet cells either in vitro or ex vivo. Adenoviruses are not only the most potent vector but also the most cytotoxic for islet cells (17). We have previously used adenoviral vectors to effectively modulate NF-κB in rat insulinoma (RIN) cell lines (18). However, the use of lentiviral vectors holds more promise for islet cell protection utilising gene therapy methods. Lentiviruses can transduce cells regardless of their mitotic state. The expression of the transgene is long-term owing to their ability to integrate into the host genome

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and also due to their low immunogenicity. Although there remain significant biosafety concerns, lentiviruses have been used to efficiently transduce islet cells both in vitro and ex vivo (19–24). Our group has previously shown the potential of lentiviral vectors to deliver shRNA to silence iNOS in the RIN cell line. The silencing of iNOS had a protective effect on β cells following cytokineinduced toxicity (25).

2. Materials 2.1. Lentiviral Vectors

1. BLOCK-iT Lentiviral RNAi expression system 2. Oligonucleotide primers encoding the three distinct iNOS shRNAs (see Note 1) 3. 1.5 mL sterile microfuge tube 4. DNase/RNase-free H2 O

2.2. Ligation

1. Ds oligos of interest: 5 nM in 1× Oligo annealing buffer 2. Ligation buffer: 50 mM Tris–HCl, 10 mM MgCl2 , 1 mM ATP, 10 mM dithiothreitol, pH 7.5, 25◦ C 3. pENTR/U6 linearised vector 4. DNase/RNase-free H2 O 5. T4 DNA ligase

2.3. Transformation

1. Competent One Shot TOP10 Escherichia coli 2. S.O.C. medium at room temperature 3. LB agar containing 50 μg/mL kanamycin pre-warmed to 37◦ C prior to use 4. Sterile toothpicks 5. Qiagen’s Miniprep kits 6. Each pENTR/U6 construct is sequenced (GATC Biotech Ltd., Cambridge, UK) to confirm the presence, correct orientation and sequence of the ds oligo insert 7. Sterile 100% glycerol

2.4. Generating Expression Clones

1. 0.5 mL microfuge tubes 2. Entry clone selected as described in Section 3.4. 3. pLenti6/BLOCK-iT – DEST vector at 150 ng/mL 4. TE buffer (pH 8.0) 5. pENTR-gus vector (50 ng/mL) 6. LR Clonase II enzyme mix 7. Proteinase K solution (20 μg/mL).

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2.5. Transforming One Shot Stbl3 Competent E.coli

1. Thawed vial of chemically competent One Shot Stbl3 cells 2. Pre-warmed S.O.C. medium 3. Pre-warmed selective plates 4. LB medium containing 100 μg/mL ampicillin

2.6. Giga Preparation of Plasmid DNA

1. Jetstar 2.0 GIGA plasmid purification kit with buffers E1–6 2. GIGA cartridges 3. Solution I: 50 mM glucose, 25 mM Tris–HCl (pH 8.0), 10 mM EDTA (pH 8.0) 4. Solution II: 0.2 M NaOH (freshly diluted from a 10 M stock), 1% SDS. 5. Solution III: 60 mL of 5 M potassium acetate, 11.5 mL glacial acetic acid, 28.5 mL H2 0. 6. Sterile 100 mL bottle 7. Isopropanol 8. 70% ethanol 9. TE buffer containing DNase-free RNase A (20 μg/mL)

2.7. Lentivirus shRNA Vectors in 293FT Cells

1. pLenti6/BLOCK-iT-DEST expression construct 2. Giga plasmid preparation kit 3. TE buffer (pH 8.0) 4. 293FT cells at 1.2 × 106 cells/mL 5. pLenti6/BLOCK-iT-DEST and the three remaining plasmids that were supplied in an optimised mixture dubbed ViraPower 6. ViraPower packaging mix 7. Serum-free Opti-MEM I Medium 8. 5 mL tubes 9. Premixed lipofectamine 2000 10. Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% foetal bovine serum 11. 100 mm diameter sterile tissue culture plates 12. 15 mL sterile, capped conical tubes

2.8. Titration of Lentiviral Stocks

1. Hela cells 2. 96 well plates 3. Blasticidin concentrations i.e., 0, 2, 4, 6, 8, 10 μg/mL 4. Trypsin (Sigma) 5. Lentiviral stocks thawed and diluted serially over the 10–2 – 10–6 range

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6. Phosphate-buffered saline 7. Crystal violet stain solution 4 g/L made in 70% ethanol (1 mL/well) 2.9. RIN-r Cell Culture

1. RIN-r cells (Gift from Steno Diabetes Centre, Gentofte, Denmark) 2. T-75 tissue culture flasks 3. RPMI 1640 supplemented with 10% (v/v) foetal bovine serum (FBS), 20 mM/L HEPES, 2 mM/L L-glutamine, 24 mM/L NaHCO3 , 100 U/mL penicillin, 100 μg/mL streptomycin 4. Trypsin 5. Coulter counter 6. Cryovials 7. Storage medium: 10% dimethyl sulfoxide (DMSO) in (DMEM) containing 10% FBS, 2 mM L-glutamine, 0.1 mM MEM non-essential amino acids, 1% penicillin/streptomycin and 1 mM MEM sodium pyruvate

2.10. Cytokine Exposure

1. Recombinant IL-1β (PromoCell, Heidelberg, Germany) at 300 and 600 U/mL 2. Cytokine cocktail (PromoCell) consisting of IL-1β, TNFα and IFN-γ at respective concentrations of 60, 185 and 14 U/mL 3. Cytokines are reconstituted in sterile PBS

2.11. Cell Viability

1. 48-well microtitre plates 2. Cell Proliferation Kit 1 which uses the MTT reagent (3-[4, 5-dimethylthiazol-2-yl]-2, 5-diphenyl tetrazolium bromide) 3. Solubilisation solution (10% w/v SDS in ultrapure water dissolved in 0.01 M HCl) 4. Microtitre plate reader

2.12. Transduction of RIN-r Cells

1. RIN-r cells 2. 6-well plates 3. Diluted lentiviral stock solution (diluted to MOI 100)

2.13. Griess Assay

1. 50 μL of cell culture medium 2. Reconstituted Griess reagent, composed of 0.1% naphthyl ethylenediamine and 1% sulphanilamide dissolved in 0.1 M HCl, 1:1 v/v. 3. BCA protein assay (Pierce)

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2.14. Western Blot Analysis

1. Whole cell lysis buffer: 5 mM Tris–HCl, 0.1 mM ethylene diamine tetraacetic acid (EDTA), 1% (w/v) sodium dodecyl sulphate (SDS) solution, 1% IGEPAL, 10% (v/v) protease inhibitor cocktail (10× stock) with pH adjusted to 7.5 2. 4% stacking/15% separation gels SDS polyacrylamide gel electrophoresis (PAGE) 3. Prestained protein markers 4. 0.2 mm nitrocellulose membrane 5. Semidry electrophoretic transfer system 6. Blocking buffer: 5% non-fat milk in PBS, 0.05% Tween 20 7. Anti-iNOS (1:2,000) is dissolved in blocking buffer 8. Anti-β-actin (1:5,000) is dissolved in blocking buffer 9. Wash buffer is composed of PBS and Tween 20 10. Sheep anti-mouse secondary antibodies conjugated to horse radish peroxidase 11. Enhanced chemiluminescence kit (ECL kit)

2.15. Detection of Cellular Oxidative Stress

1. 21 gauge and 26 gauge needles 2. Reaction mixture: 100 μL 1 M Tris–HCl, 20 μL 0.1 M γ-L-Glutamyl-L-cysteinyl-glycine (GSH), 100 μL glutathione (10 U/mL), 100 μL 2 mM NADPH, 610 μL ultrapure H2 O 3. Tert-butylhydroperoxide dissolved 1:1,000, v/v in water

3. Methods In order to produce the appropriate shRNAs specific for iNOS RNAi, this investigation employed the BLOCK-iT Lentiviral RNAi Expression System to facilitate the generation of a replication-incompetent lentivirus. The kit employs Gateway Technology which encompasses a universal cloning strategy that exploits the site-specific recombination attributes of bacteriophage λ. The technique allows for the rapid and efficient movement of a DNA sequence of interest into multiple compatible vector systems which ultimately permit the expression of shRNAs of interest in mammalian cells. The concept involves the following steps: double-stranded oligonucleotides are first designed to encode the shRNA of interest; these oligos are then cloned into a suitable intermediate vector, pENTR/U6 to create an entry clone; the expression clone is next generated by recombining the

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entry clone with the pLenti6/Block-iT – DEST vector; the resultant expression clone is then cotransfected into an appropriate packaging cell line (i.e. 293FT cells) for vector particle assembly and production. The lentiviral vector is then titred and used for mammalian cell transduction using the standard respective methodologies. Titration of the lentiviral stocks was necessary to control the number of integrated copies of the vectors into the target cells and to facilitate reproducible gene knockdown results. To determine the lentiviral vector stock titres, blasticidin resistance in stably transduced Hela cells was analysed. As noted in our introduction the induction of iNOS and the subsequent production of NO in β cells have been implicated as a critical mediator of β cell death following cytokine exposure. The Griess assay is one of the most popular and simplest methods used to detect nitrite accumulation. In order to evaluate the extent of nitric oxide production induced by cytokine exposure and subsequent potential knockdown of expression by iNOS shRNA, media samples from RIN-r cells cultured in vitro were analysed for accumulated nitrite. 3.1. Lentiviral Vectors Expressing Rat iNOS Specific shRNAs

1. Design the oligonucleotide primers encoding the three distinct iNOS shRNAs according to the specified guidelines by Ambion (see Note 2). The DNA oligonucleotides encoding the shRNAs should be denoted NOS1, NOS2 and NOS3 (Fig. 12.1). 2. Anneal the primers as follows: Incubate 5 μL of both the top strand and bottom strand DNA oligos (at 200 μM) in a 1.5 mL sterile microfuge tube with 2 μL 10× oligo annealing buffer and 8 μL DNase/RNase-free H2 O at 95◦ C for 4 min. Following this, remove the tube and cool at room

Oligonucleotide sequences designed specific for rat iNOS gene knockdown NOS1 DsOligo

NOS2 DsOligo

NOS3 DsOligo

Fig. 12.1. Oligonucleotide sequences designed specific for rat iNOS gene knockdown. Three distinct sequences were designed and denoted NOS1, NOS2 and NOS3. Once annealed the ds oligonucleotide sequences encode three distinct shRNAs of interest.

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temperature for 5–10 min. Centrifuge the samples briefly and gently vortex. 3. Dilute (100-fold) the annealing mixture (20 μL) to a working concentration of 500 nM in DNase/RNase-free H2 O. Dilute this solution further (100-fold) to 5 nM in 1× oligo annealing buffer. 4. Label the ds oligos and store at –20◦ C. The 50 mM, 500 nM and 5 nM ds oligo stocks can be used for preparation of new diluted ds oligo when needed, gel analysis to check the integrity of annealed oligos and to clone into the pENTR/U6 plasmid, respectively. 3.2. Ligation Reaction into the pENTR/U6

1. Thaw the ds oligos of interest (i.e. 5 nM in 1× oligo annealing buffer) on ice before use. 2. Add 5× ligation buffer (4 μL), 2 μL of 0.5 ng/mL pENTR/U6 linearised vector, 1 μL of ds oligo, 12 μL DNase/RNase-free H2 O and 1 μL of T4 DNA ligase (1 U/μL). 3. Mix the ligation reaction by pipetting up and down and incubate at room temperature for 20 min. Place the reaction mixture on ice before transforming the One Shot TOP10 competent E.coli.

3.3. Transforming the One Shot TOP10 E. coli

1. Add 2 μL of the ligation reaction into a vial of competent E. coli and mix gently by flicking the tube. Incubate the mixture on ice for 10 min before heat shocking the cells at 42◦ C for 30 s by using a heating block without shaking. 2. Transfer the tubes immediately onto ice and add 250 μL of room temperature S.O.C medium. Shake the tubes at 200 rpm at 37◦ C for 1 h (see Note 3). 3. Spread 20 and 200 μL aliquots from each transformation aseptically onto pre-warmed LB agar plates containing 50 μg/mL kanamycin. Invert the plates and incubate overnight at 37◦ C. 4. The following day pick five colonies from each ligation mixture and analyse the transformants.

3.4. Analysing Transformants

1. Pick kanamycin-resistant clones and culture overnight in LB medium, supplemented with 50 μg/mL kanamycin. 2. Using Qiagen’s Mini Prep kits isolate plasmid DNA. Each pENTR/U6 construct is sequenced (GATC Biotech Ltd.) to confirm the presence, sequence and correct orientation of the ds oligo insert. To facilitate sequence analysis of the pENTR/U6 clones, use the U6 forward and M13 reverse primers.

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3. Following identification of correct entry clones, streak the original colonies on an LB plate containing kanamycin to obtain single colonies. Isolate single colonies and inoculate into 3 mL of LB supplemented with 50 μg/mL kanamycin. Grow the culture overnight. 4. Using cryovials, add 0.85 mL aliquots of the bacterial culture to 0.15 mL 100% glycerol and store at –80◦ C. 5. Following confirmation of the pENTR/U6 clones, perform a recombination reaction between the pENTR/U6 construct and the destination vector to generate an expression clone. 3.5. Expression Clones

1. Add the following components to a 0.5 mL microfuge tube at room temperature: 100 ng of the entry clone, 1 mL pLenti6/BLOCK-iT – DEST vector (150 ng/mL), TE buffer (pH 8.0) up to 8 mL. 2. As a positive control for the reaction, substitute 2 mL of the pENTR-gus vector (50 ng/mL) for the entry clone. 3. Remove the LR Clonase II enzyme mix from –20◦ C and thaw on ice for 2–3 min. Vortex the mixture twice for 2–3 s before adding 2 mL of LR Clonase II enzyme mix. Vortex and incubate the reaction mixture at 25◦ C for 1 h. 4. Add 1 μL of proteinase K solution (20 μg/mL) to each reaction mixture and incubate for 10 min at 37◦ C. Transform competent One Shot Stbl3 E.coli cells with the reaction mixture.

3.6. Transforming One Shot Stbl3 Competent E. coli

1. Add 2–3 μL of the LR recombination reaction to one thawed vial of chemically competent One Shot Stbl3 cells. Tap the vial to ensure thorough but gentle mixing of the reaction components. 2. Place the vial on ice for 30 min and then heat shock the cells for 45 s at 42◦ C using a heat block without shaking. Remove the vial and place on ice for 2 min before adding of 250 μL pre-warmed S.O.C. medium to each vial. 3. Seal the vial and shake horizontally at 37◦ C for 1 h at 225 rpm. Spread aliquots of the transformation mixture (20 and 200 μL) on pre-warmed selective plates. The remaining mix can be stored short term at 4◦ C. 4. It is not necessary to sequence the expression clones as transfer of the U6 RNAi cassette from pENTR/U6 into the pLenti6/Block-iT-DEST vector preserves orientation of the cassette. Once generated, maintain the expression clones in LB medium containing 100 μg/mL ampicillin.

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3.7. Giga Preparation of Plasmid DNA

1. Use the Jetstar 2.0 GIGA plasmid purification kit to amplify large-scale preparations of plasmid DNA. 2. Centrifuge E.coli bacterial cells at 4,000×g for 15 min and aspirate the supernatant. 3. Resuspend the bacterial pellets in 125 mL buffer E1 until there are no cell clumps present. 4. Add 125 mL buffer E2 to lyse the cells, mix gently and thoroughly, incubate the mixture at room temperature for 5 min. 5. Add 125 mL buffer E3 and followed by gentle but thorough mixing to stop cell lysis. 6. Centrifuge the mixture at 12,000×g for 20 min at room temperature. Decant the supernatant into a sterile container. 7. Using a vacuum, equilibrate the GIGA cartridges by passing 200 mL equilibration buffer E4 through the cartridge. 8. Discard the flow-through and load the cleared lysate onto the filter cartridge. Use a vacuum to pass the lysate through the resin. Wash the resin twice with 300 mL of buffer E5 before eluting the plasmid. 9. Attach the filter cartridge to a sterile 100 mL bottle with a 45 mm neck. Apply 100 mL of elution buffer E6 into the cartridge. Use the vacuum briefly to initiate the flow of elution buffer containing the plasmid DNA. 10. Precipitate the plasmid DNA with 70% isopropanol and centrifuge at 15,000×g for 30 min at 4◦ C. Wash the precipitated DNA with 70% ethanol and again centrifuge at 15,000×g for 5 min at 4◦ C. Aspirate the supernatant and dry the pellet at room temperature for 10 min before redissolving the DNA in a suitable volume of sterile TE containing DNase-free RNase A (20 μg/mL).

3.8. Production of Lentivirus shRNA Vectors in 293FT Cells

1. Amplify the pLenti6/BLOCK-iT-DEST expression construct using the Jetstar 2.0 Giga plasmid purification kit. 2. Resuspend the purified pLenti6/BLOCK-iT plasmid in sterile TE buffer (pH8.0) to a concentration of 1 mg/mL. 3. Cotransfect the 293FT cells with the four plasmid set, i.e., the pLenti6/BLOCK-iT-DEST, pLP1, pLP2 and pLP/VSV-g plasmids. 4. Prepare the transfection complexes as follows: a. Combine 9 μg of ViraPower packaging mix with 3 μg of pLenti6/BLOCK-iT-DEST expression plasmid and dilute in 1.5 mL serum-free Opti-MEM I Medium

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b. In a separate 5 mL tube, add 36 μL of premixed lipofectamine 2000 with 1.5 mL of serum-free Opti-MEM and incubate at room temperature for 5 min. c. Combine the lipofectamine 2000 and DNA complexes and incubate at room temperature for 20 min. 5. Trypsinise the 293FT cells, count and resuspend in serum supplemented medium at 1.2 × 106 cells/mL. 6. Add the DNA-lipofectamine 2000 complex to a 100 mm diameter sterile tissue culture plate containing 5 mL of growth medium without antibiotics. 7. Add 6 ×106 cells to each plate, mixed gently and incubate overnight at 37◦ C in 5% CO2 /95% air. Remove the media the following day and replaced with complete DMEM (DMEM containing 10% FBS, 2 mM L-Glutamine, 0.1 mM MEM non-essential amino acids, 1% penicillin/streptomycin and 1 mM MEM sodium pyruvate). 8. 60 h post-transfection harvest the virus-containing supernatants into 15 mL sterile capped conical tubes. Centrifuge the tubes at 3,000 rpm for 5 min at 4◦ C to pellet cell debris. 9. Aliquot (1 mL) the viral supernatants into cryovials. Store the stocks at –80◦ C. 3.9. Titration of Lentiviral Stocks

1. Plate Hela cells in 96-well plates at 1,000 cells per well and allowed to attach for 24 h. 2. To determine dose response, expose the cells to various concentrations of blasticidin (0, 2, 4, 6, 8, 10 μg/mL). Replenish the selective medium every 3–4 days (see Note 4). The concentration of blasticidin that killed the cells after 10 days was determined to be 4 μg/mL. 3. 24 h prior to transduction, trypsinise fresh Hela cells, count and seed in a 6-well plate at a cell density of 1.5 × 105 cells/well. Allow cells to attach for 24 h. 4. Thaw aliquots of lentiviral stocks and serially diluted over the 10–2 –10–6 range. Make all dilutions in complete culture medium to a final volume of 1 mL. 5. Aspirate the culture medium of Hela cells and replace with 1 mL of the lentiviral dilution. 6. Gently rock the plates to ensure even distribution of the lentiviral solution. Incubate overnight at 37◦ C. Remove the lentiviral solutions by aspiration and replace with complete culture medium. Allow cells to recover from transduction for a further 24 h before the addition of blasticidin-supplemented media to select for stably transduced cells.

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7. Replenish blasticidin-supplemented media every 3–4 days. Following 10 days of blasticidin exposure, remove culture medium and wash twice with warm PBS. 8. Add 1 mL/well of crystal violet stain and incubated at room temperature for 10 min. Remove excess stain and wash with PBS before counting the colonies. Determine the titre of the lentiviral stock by multiplying the number of positively stained colonies by the relevant dilution. Titres are expressed in transduction units/mL. 3.10. RIN-r Cell Culture

1. Culture RIN-r cells at 37◦ C in T-75 tissue culture flasks in RPMI 1640 supplemented with 10% (v/v) FBS, 20 mM/L HEPES, 2 mM/L L-glutamine, 24 mM/L NaHCO3, 100 U/mL penicillin, 100 μg/mL streptomycin under a humidified atmosphere of 5% CO2 /95% Air. 2. Passage cells every 3–4 days following trypsinisation. Obtain accurate cell counts using a Coulter counter to ensure precise plating densities. 3. Store cells stocks in 10% DMSO/90% FBS for long-term storage in liquid N2 . Prior to reuse remove DMSO by centrifugation and resuspend the cell pellet in complete culture medium.

3.11. Cytokine Exposure

1. Seed RIN-r cells in a 48-well microtitre plate at a density of 2.5 × 104 cells/well in a final volume of 300 μL/well. 2. Allow the RIN-r cells to recover for 24 h after plating in 48 well plates. 3. Expose the RIN-r cells either to IL-1β alone, at 0.5× (i.e. 300 U/mL) and 1× (i.e. 600 U/mL) or to a 0.5× and 1× cytokine cocktail (1× cytokine cocktail consists of IL1β, TNF-α and IFN-γ at respective concentrations of 60, 185 and 14 U/mL). 4. Expose to the cytokines for periods of 24, 48, 72 and 120 h (Fig. 12.2). Assess cell viability as outlined below.

3.12. Cell Viability

1. Determine the cell viability of the RIN-r cells using the MTT reagent (3-[4, 5-dimethylthiazol-2-yl]-2, 5-diphenyl tetrazolium bromide) and the MTT cell proliferation kit 1 assay (Roche). 2. Following the cytokine incubation period, 0.5 mg/mL MTT labelling reagent was added to each well. 3. Incubate the microtitre plates at 37◦ C, 5%, CO2 , 95% air for 4 h before adding 200 μL of solubilisation solution (10% SDS in 0.01 M HCl) to each well.

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*

* * **

Fig. 12.2. Establishing a model for the induction of iNOS in RIN-r cells exposed to 100 U/mL IL-1β. IL-1β induces a time-dependent accumulation of nitrite when compared to baseline levels of nitrite in the media of RIN-r cells. All data are means ± SD. Experiments were performed in triplicate with an n = 4. ∗ Significance established at p < 0.005. ∗∗ Significance established at p < 0.0005. All significance established relative to non-cytokine-exposed RIN-r cells.

4. Incubate the microtitre plates at 37◦ C, 5%, CO2 , 95% air overnight, after which, record the A550 nm using a plate reader (SpectraMax). 3.13. Transduction of RIN-r Cells and Analysis

1. Plate RIN-r cells in 6-well plates at a density of 3 × 105 cells/well and allow to attach for 24 h before transduction with the diluted lentiviral stock solution (diluted to MOI 100). 2. Use the minimal volume of fresh medium required to cover the cell monolayer as a diluent (750 μL for 6-well plates and 100 μL for 48-well plates) (see Note 5). 3. Incubate the plates at 37◦ C overnight. Replace the viruscontaining medium with complete culture medium (see Note 6). 4. Leave the plates for 3 days and then expose the RIN-r cells to the appropriate concentrations of IL-1β for the appropriate time period. Collect the Rin-r cell medium and lysates for nitrite quantification and iNOS protein expression following cytokine exposure.

3.14. Griess Assay

1. Incubate 50 μL of cell culture media with reconstituted Griess reagent (1:1) (0.1% naphthyl ethylenediamine and 1% sulphanilamide in 0.1 M/L HCl, 1:1 vol/vol) for 15 min in the dark at room temperature. Perform a nitrite standard curve to determine the nitrite concentrations in each media sample. 2. To normalise nitrite concentration to cell numbers plated, harvest the cells, lyse and assay for protein using the Brad-

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ford protein assay. Determine protein concentrations at A540 nm using a plate reader (SpectraMax). 3. Normalise nitrite concentrations to total protein concentrations determined for each sample to correct for discrepancies in cell seeding density. This is expressed in micromoles (μM) of accumulated nitrite/milligram (mg) of total protein. 3.15. Western Blot Analysis

1. Harvest the cells by scraping and centrifuge at 2,300 rcf for 5 min. 2. Extract the total protein from the cell pellets via resuspension in 110 μL of whole cell lysis buffer (5 mM Tris–HCl, 0.1 mM EDTA, 1% (w/v) SDS, 1% IGEPAL, 10% (v/v) protease inhibitor cocktail, pH 7.5). Place on ice for 20 min. 3. Centrifuge at 2,300 rcf for 5 min. Collect the supernatant and determine the total protein via the Bradford assay (Pierce). 4. Load prestained protein markers and 15 μg of protein per well on a 4% stacking/15% for separation SDS-PAGE gel. 5. Transfer resolved proteins to a 0.2 mm nitrocellulose membrane using a semidry electrophoretic transfer system for Western blot analysis. 6. Place membranes in blocking buffer (5% non-fat powdered milk in PBS/0.05% Tween 20) and shake overnight at 4◦ C. 7. Incubate the membranes with primary anti-iNOS (1:2,000), and anti-β-actin (1:5,000) antibodies in blocking buffer overnight at 4◦ C. 8. Wash the membranes three times in PBS-Tween. Incubate the membrane for 1 h at room temperature with sheep antimouse secondary antibodies conjugated to HRP. 9. Detect protein bands using an enhanced chemiluminescence kit (ECL kit) followed by exposure to x-ray film. β-Actin served as a housekeeping gene.

3.16. Detection of Cellular Oxidative Stress

1. Harvest the Rin-r cells by scraping and centrifuge at 2,300 rcf for 5 min. Lyse cells by slowly passing through 21 gauge and 26 gauge needles consecutively 5–6 times on ice. 2. Collect the supernatant, use 50 μL aliquots from the sample lysate to determine the subsequent glutathione peroxidase (GPx) activity. 3. Incubate the samples in a reaction mixture (100 μL 1 M Tris–HCl, 20 μL 0.1 M GSH, 100 μL glutathione (10 U/mL), 100 μL 2 mM NADPH, 610 μL ddH2 O) for 10 min at 37◦ C. 4. Add 20 μL of t-butylhydroperoxide (1:1,000, v/v) to the reaction mixture and incubate for a further 10 min at 37◦ C.

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Read the absorbance against distilled water at A340 nm . Blank each sample was appropriately using the reaction mixture without GSH. Calculate GPx activity using the formula: OD (blank– system) × dilution factor to 1 mL/mg cell protein × 62.2 = U/mg cell protein.

4. Notes 1. NOS shRNAs were designed according to specified guidelines by Ambion. 2. The primers were commercially synthesised by Invitrogen and annealed to create a double-stranded oligonucleotide. 3. The tubes should be tightly capped, covered in foil to minimize condensate buildup on the cap undersurface and shaken at 200 rpm at 37◦ C for 1 h. 4. Care should be taken to ensure gentle aspiration and therefore reduce the possibility of inadvertently aspirating loosely adhered cells. 5. In an effort to maximise transduction efficiency the minimal volume of fresh medium required to cover the cell monolayer is used as diluent. 6. To increase viral coverage the plates need to be rocked gently back and forth once every 10 s for 2 min before placing them at 37◦ C overnight. References 1. Debray-Sachs, M., Carnaud, C., Boitard, C., Cohen, H., Gresser, I., Bedossa, P., et al. (1991) Prevention of diabetes in NOD mice treated with antibody to murine IFN gamma. J Autoimmun 4, 237–248. 2. Larsen, J. L., Duckworth, W. C., Stratta, R. J. (1994) Pancreas transplantation for type I diabetes mellitus. Do the benefits offset the risks and cost?. Postgrad Med 96, 105–111. 3. Ryan, E. A., Paty, B. W., Senior, P. A., Shapiro, A. M. (2004) Risks and side effects of islet transplantation. Curr Diab Rep 4, 304–309. 4. Reckard, C. R., Ziegler, M. M., Barker, C. F. (1973) Physiological and immunological consequences of transplanting isolated pancreatic islets. Surgery 74, 91–99. 5. Shapiro, A. M., Lakey, J. R., Ryan, E. A., Korbutt, G. S., Toth, E., Warnock, G. L., et al. (2000) Islet transplantation in seven

patients with type 1 diabetes mellitus using a glucocorticoid-free immunosuppressive regimen. N Engl J Med 343, 230–238. 6. Balamurugan, A. N., Bottino, R., Giannoukakis, N., Smetanka, C. (2006) Prospective and challenges of islet transplantation for the therapy of autoimmune diabetes. Pancreas 32, 231–243. 7. Rabinovitch, A., Sorensen, O., SuarezPinzon, W. L., Power, R. F., Rajotte, R. V., Bleackley, R. C. (1994) Analysis of cytokine mRNA expression in syngeneic islet grafts of NOD mice: interleukin 2 and interferon gamma mRNA expression correlate with graft rejection and interleukin 10 with graft survival. Diabetologia 37, 833–837. 8. Corbett, J. A., McDaniel, M. L. (1995) Intraislet release of interleukin 1 inhibits beta cell function by inducing beta cell expression

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of multiple chemokines and chemokine receptors in murine beta cells and pancreatic islets. Am J Transplant 3, 1230–1241. McCabe, C., Samali, A., O‘Brien, T. (2006) beta cell cytoprotective strategies: establishing the relative roles for iNOS and ROS. Biochem Biophys Res Commun 342, 1240–1248. Ju, Q., Edelstein, D., Brendel, M. D., Brandhorst, D., Brandhorst, H., Bretzel, R. G., et al. (1998) Transduction of non-dividing adult human pancreatic beta cells by an integrating lentiviral vector. Diabetologia 41, 736–739. Gallichan, W. S., Kafri, T., Krahl, T., Verma, I. M., Sarvetnick, N. (1998) Lentivirusmediated transduction of islet grafts with interleukin 4 results in sustained gene expression and protection from insulitis. Hum Gene Therapy 9, 2717–2726. Lu, Y., Dang, H., Middleton, B., Zhang, Z., Washburn, L., Campbell-Thompson, M., et al. (2004) Bioluminescent monitoring of islet graft survival after transplantation. Mol Therapy 9, 428–435. Giannoukakis, N., Mi, Z., Gambotto, A., Eramo, A., Ricordi, C., Trucco, M., et al. (1999) Infection of intact human islets by a lentiviral vector. Gene Therapy 6, 1545–1551. Kobinger, G. P., Deng, S., Louboutin, J. P., Vatamaniuk, M., Matschinsky, F., Markmann, J. F. (2004) Transduction of human islets with pseudotyped lentiviral vectors. Hum Gene Therapy 15, 211–219. Okitsu, T., Kobayashi, N., Totsugawa, T., Maruyama, M., Noguchi, H., Watanabe, T., et al. (2003) Lentiviral vector mediated gene delivery into non-dividing isolated islet cells. Transplant Proc 35, 483. McCabe, C.„ O‘Brien, T. (2007) The rational design of beta cell cytoprotective gene transfer strategies: targeting deleterious iNOS expression. Mol Biotechnol 37, 38–47.

Chapter 13 Characterization of Nitric Oxide Delivery Systems Produced By Various Nanotechnologies Chi H. Lee Abstract Advanced formulations having the capability of long-term protection and preserving the integrity of the nitric oxide molecule were developed for the efficient delivery of nitric oxide. The methods for the production and delivery of nitric oxide produced from diethylenetriamine diazeniumdiolate or Snitrosoglutathione as a nitric oxide donor to desired target sites in a controlled and sustained release manner are described. The pharmacological assessments of advanced formulations using in vitro and in vivo tests are also described in detail. These systems can be used for the treatment of cardiovascular diseases and sexual dysfunctions. Key words: Nitric oxide delivery, PLGA microparticle, polymeric GSNO film, cell cytotoxicity, blood flow.

1. Introduction Nitric oxide (NO) as a neurotransmitter plays integral physiological roles including the blood flow regulation, immune response, and wound healing process (1, 2). NO produced from L-arginine diffuses from endothelial cells into smooth muscle, causing an increase of cGMP and vasodilation of vessels (3, 4). When this process is inadequate, the arterial dilation process is hindered, resulting in a reduced blood flow (5). The endothelial dysfunction subsequently causes smooth muscle proliferation, platelet and monocyte adhesiveness enhancement, and unbalance in the blood flow regulation, leading to various cardiovascular diseases including atherosclerosis, hypertension, and sexual abnormalities (1, 2, 5). H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_13, © Springer Science+Business Media, LLC 2011

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Considering wide applications of NO in the physiological process, the delivery of exogenous NO will produce numerous beneficial effects and can be used to prevent or treat various diseases (6). Consequently, the development of suitable and advanced systems intended for efficient NO delivery is integral in optimizing its effects and maximizing its benefits. Advanced formulations, such as microparticles and polymeric films, containing diethylenetriamine diazeniumdiolate (DETA NONOate) or S-nitrosoglutathione (GSNO) as a NO donor were developed to take advantage of their versatility including high loading efficacy and biocompatibility (7, 8). Further evaluation processes of NOreleasing formulations using in vitro and in vivo tests were followed to validate their pharmacological effects (9). Those tests include the assessments of entrapment efficacy, morphology, stability, NO release profiles, cytotoxicity, and in vivo blood flow changes.

2. Materials 2.1. Property of NO

2.2. Reagents and Solvents

NO is a colorless gas and reacts readily with oxygen to form brown nitrogen dioxide. Under physiological conditions, NO can be transformed with distinctive chemical reactions to different redox forms, such as NO, NO+ , and NO– . The biological halflife of NO is very short and it is readily soluble in water, showing about 4.7 parts per 100 parts volume at 20◦ C (1 atm). NO has a melting point of –163◦ C, boiling point of –151.7◦ C, and liquid density of 1.269 (at boiling point). NO is a powerful endogenous vasodilator (i.e., endothelial-derived relaxing factor) as well as a neurotransmitter, acting as a signaling molecule in the body. 1. PLGA 5050 (I.V. = 0.26 dL/g) and PLGA 7525 (I.V. = 0.27 dL/g) Lakeshore Biomaterials (see Note 1) 2. Diethylenetriamine diazeniumdiolate (DETA NONOate) (see Note 2) 3. S-Nitrosoglutathione (GSNO) (see Note 2) 4. Poly(vinyl alcohol) (PVA) 5. Light mineral oil 6. Span80 7. Carbopol 934P 8. Hydroxypropyl methylcellulose (HPMC, Methocel E4M premium) 9. Polyethylene glycol (PEG, MW 1000) 10. Glutathione (γ-Glu-Cys-Glu, GSH)

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11. Sodium nitrite (NaNO2 ) (see Note 3) 12. Porcine aortic valve interstitial cells (PAVICs) 13. Normal vaginal epithelial cells (VK2E6E7) 14. Cell titer 96 aqueous cell proliferation assay solution 15. Citric acid 16. Sodium citrate 17. Phosphate-buffered saline 18. Sodium hydroxide (NaOH) 19. Hexane

3. Methods As NO has a very short half-life (3–4 s), the target specific delivery of NO with a controlled release rate has been a major challenge in the development of a topical delivery system for NO. To date, NONOate and GSNO have been widely used as NO donors, since they spontaneously release two equivalents of NO at a reliable first-order rate through the hydrolysis process (10). The decomposition of DETA NONOate or GSNO is nearly instantaneous at pH of 5.0 or less. Therefore, the NO donor should be stabilized under acidic environment for a prolonged delivery of NO. Poly(D,L-lactic acid-co-glycolic acid) (PLGA) microparticles and polymeric films (Carbopol, HPMC and PEG) were developed as a carrier vehicle for DETA NONOate and GSNO, respectively, considering that PLGA or polymer combinations are biocompatible and widely used as biodegradable polymers (7, 8). Polymers would prevent DETA NONOate or GSNO from decomposition at acidic pH, releasing NO to the target sites in a controlled manner. Depending on the target sites and topical mucoadhesivity, microparticles or films can be selected as NO carriers. It was reported that topically administered NO donor increased the blood flow without affecting cardiovascular functions, implying NO therapy would be suitable for the treatment of cardiovascular diseases and female sexual arousal disorder (FSAD) (9, 11). 3.1. Preparations of Microparticles

3.1.1. Single Emulsion Methods

PLGA microparticles were fabricated by single or double emulsion solvent evaporation methods (12). The procedure for each method is as follows. 1. Prepare the PLGA polymer solution (5% (w/v)) in dichloromethane (DCM). 2. Disperse the NO donors (5% (w/w) of DETA NONOate) in PLGA polymer solutions. Place on an ice bath and probe sonicate at 20 W for 20 s.

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3. Add drug-dispersed polymer solution to the PVA solution (2% (w/v)) to form a water in oil (w/o) emulsion. 4. Add the drug-dispersed polymer solution to light mineral oil containing 2% (w/v) of the Span80 to form an oil in oil (o/o) emulsion. 5. Microparticles produced by single emulsion methods were spherical with sizes ranging from 19 to 40 μm. 6. Entrapment efficiency of DETA NONOate in microparticles produced by the single emulsion method was low (4–5.2%) as compared with those produced by double emulsion methods (see Note 4). 3.1.2. Double Emulsion Methods

1. Dissolve 2.5, 5, and 10% (w/w) of DETA NONOate in 10 mM aqueous NaOH solution to minimize the decomposition of DETA NONOate (see Note 5). 2. Pour the drug solution into PLGA 5050 or 7525 solutions (5% (w/v)). 3. Form a primary emulsion (w/o) by probe sonication at 20 W for 20 s on an ice bath. 4. Add the primary emulsion in a dropwise manner to aqueous PVA solutions (2% (w/v)), PVA solutions contain 5 mM NaOH (i.e., an alkalic compound) or light mineral oil with Span80 to form water in oil in water (w/o/w), water in oil in alkaline water (w/o/wa ), and water in oil in oil (w/o/o) emulsion, respectively (see Note 6). 5. Stir the final emulsion at 450 rpm at 25◦ C in a fume hood for 5 h to remove the organic solvent. 6. Recover the microparticles by washing three times with deionized water and hexane. 7. Filter the recovered microparticles with a 0.2 μm cut-off membrane to remove excess PVA or light mineral oil with Span80 (see Note 7). 8. Harvest the obtained microparticles for 24 h by freezedrying and subject to morphology tests. Store microparticles at –20◦ C. 9. Use a scanning electron microscopy (SEM; FEG ESEM XL 30, Hillsboro, OR) to examine the shape and surface morphology of the microparticles. 10. Mount a small amount of the microparticles on doublesided tape attached to a metallic sample stand. Spray-coat the microparticles with gold palladium at 0.6 kV prior to SEM imaging. Microparticles from all formulations displayed spherical shapes and there were no obvious differences in shape among the formulations prepared by three different emulsion methods

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(Fig. 13.1s). Microparticles made by the w/o/w method had highly porous surface, whereas w/o/o significantly reduced pores on the microparticles. Microparticles made by w/o/wa method

Fig. 13.1. The results of the SEM study on microparticles. a Microparticles made by w/o/w method; PLGA 5050 (left) and PLGA 7525 (right), b Microparticles made by w/o/o method; PLGA 5050 (left) and PLGA 7525 (right), c Microparticles made by w/o/wa method; PLAG 5050 (left) and PLGA 7525 (right), d Internal morphology of the micropaticle (PLGA 5050, w/o/wa ) (Cited from reference (7).

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significantly reduced porosity as compared with those made by w/o/w. 3.2. Preparation of the S-Nitrosoglutathione (GSNO) Film

1. Prepare the GSNO films made of Carbopol, HPMC, and PEG using the solvent evaporation method (13). 2. Add the carbopol–HPMC mixture with the PEG to produce the final polymeric solution (Cabopol:HPMC:PEG = 1.5:1.5:1). 3. Stir the 6.7% w/v polymeric solution overnight and keep at room temperature until all the air bubbles are removed (see Note 8). 4. Add various amounts (5–30 wt.%) of GSNO to the polymer solution. 5. Stir the solution for 30 min at room temperature, then sonicate to remove bubbles. 6. Measure the viscosity of the polymer-GSNO solutions using a cone and plate viscometer (Thomas Scientific, Swedesboro, NJ) at 37◦ C. 7. Cast the polymer-GSNO solution in a petri dish and dry at room temperature. 8. Examine the surface morphology of the GSNO film by SEM. 9. Mount the samples on double-sided tape and spray-coated with gold palladium at 0.6 kV. 10. Peel off the films from the tape and examine under SEM or store at –20◦ C until required (see Note 9). 11. An example of the GSNO films inspected by scanning electron microscope (SEM) (Fig. 13.2) (a) blank film (no GSNO) and (b) GSNO film (20 wt.% loading dose). Although the film with 20 wt.% loading dose of GSNO had slightly rougher surface as compared with the blank film, it still had homogenous surface with no pores.

(a)

(b)

Fig. 13.2. Surface morphology of (a) blank film (no GSNO) and (b) GSNO film (20 wt.% loading dose) obtained by scanning electron microscope (SEM) (Cited from reference 8).

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3.3. Evaluation of Microparticles 3.3.1. AFM Study on Morphology of PLGA Microparticles

1. Examine the size and morphology of the PLGA microparticles by atomic force microscopy (AFM) to provide detailed information on their spherical morphology and geometry (14). 2. Use an AFM equipped with a Digital Nanoscope IV Bioscope (Veeco Instruments, Santa Barbara, CA, USA) and a microscope with a carbon nanotube (CNT) tip (see Note 10). 3. Perform the AFM in the tapping mode with the conditions set at a resonance frequency of 220 kHz and a nominal force constant of 36 N/m. Set the scan speed according to the scan size with a scan frequency from 0.5 to 1.5 Hz. 4. Obtain images to estimate particle size distribution by a curvature reconstruction method (CRM), displaying amplitude, height and phase signal of the cantilever in the trace direction. The results of this study demonstrated that variance of the microparticle size was within an acceptable range and surface morphology of PLGA microparticles was found to be smooth with slight aggregation, whose degree varying according to the manufacturing methods.

3.3.2. Assessment of Particle Size Distribution of PLGA Microparticles

1. Determine particle size microscopically at the magnification of ×20 (Carl Zeiss, Germany) with the aid of a stage and eye piece micrometer (15). 2. Mount a random sample of microparticles on a glass slide with a drop of distilled water. 3. Use at least 200 particles of each batch for the assessment of the particle size distribution (PSD). The microparticles made by double emulsion methods (w/o/w, w/o/wa and w/o/o) had a greater particle size than those made by single emulsion methods (o/w and o/o) as shown in Table 13.1. The microparticles prepared by w/o/o methods had a particle size around 40 μm, whereas w/o/w methods produced microparticles with a large particle size (68–83 μm).

3.3.3. Assessment of Entrapment Efficiency of the NO Donors in PLGA Microparticles

1. Determine the content of DETA NONOate in microparticles by PLGA digestion (12). 2. Suspend a known amount of the microparticle in 1 M NaOH and sonicate in a bath for 10 min. 3. Stir the suspension to completely hydrolyze PLGA. 4. Measure the amount of NO in the final transparent hydrolysate using a spectrophotometer at a wavelength of 252 nm (15).

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Table 13.1 Properties of DETA NONOate-loaded PLGA microparticles (Cited from reference 7) Particle sizeb (μm)

Entrapment efficiencya (%)

Loading content (mg)

Method

Yielda

5050c O/W

87.0 ± 6.6

23 ± 11

4.0 ± 0.8

0.2 ± 0.04

5050 O/O

95.3 ± 3.0

19 ± 10

5.2 ± 1.5

0.3 ± 0.08

5050 W/O/W

92.6 ± 4.8

83 ± 23

16.7 ± 2.2

0.8 ± 0.11

7525d W/O/W

86.3 ± 4.5

68 ± 20

18.2 ± 3.6

0.9 ± 0.18

5050 W/O/O

96.1 ± 4.6

38 ± 19

30.7 ± 2.8

1.5 ± 0.14

(%)

7525 W/O/O

95.5 ± 3.8

41 ± 21

34.2 ± 2.4

1.7 ± 0.12

5050 W/O/Wa

90.1 ± 4.6

78 ± 23

72.7 ± 2.3

3.6 ± 0.12

7525 W/O/Wa

94.5 ± 3.8

81 ± 26

73.2 ± 3.5

3.7 ± 0.18

Loading dose of DETA NONOate was 5 mg (5% w/w) for all formulations. an = 3 b n ≥ 200 c 5050 = PLGA 5050 d 7525 = PLGA 7525

5. Generate a calibration curve using the standard solutions of five different concentrations of DETA NONOate by mixing blank PLGA microparticles and known amounts of DETA NONOate in 1 M NaOH. 6. Calculate the entrapment efficiency (%) of the samples prepared in triplicate using the following equation:

Entrapment efficiency (%) =

Amount of remaining DETA NONOate in the microparticles Amount of initially added DETA NONOate

× 100

When single emulsion methods were used, entrapment efficiency was very low (4–5.2%). Double emulsion methods (w/o/w and w/o/o) enhanced entrapment efficiency up to 18.2 and 34.2%, respectively. Double emulsion methods (w/o/wa ), in which the alkaline solution was used as a dispersing phase, enhanced entrapment efficiency up to 73.2% that is significantly higher than those obtained by other double emulsion methods (w/o/w and w/o/o). 3.3.4. Assessment of Stability of the NO Donors in the Microparticles

1. Maintain microparticles prepared by w/o/wa method (i.e., produced the microparticles with the most stable and entrapment efficiency) in a temperature-humidity chamber (Hotpack, Philadelphia, PA, USA) at 25◦ C with the relative humidity of 60% or for long-term storage freeze at –20◦ C (see Note 11).

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2. After a predetermined time interval digest triplicate sets of the microparticles using 1 M NaOH. 3. Analyze the remaining intact DETA NONOate by spectrophotometer at a wavelength of 252 nm. DETA NONOate in the microparticles remained intact at –20◦ C for at least 1 month. At room temperature DETA NONOate in the microparticles is rapidly decomposed. About 50% of DETA NONOate was decomposed in 5 days and only 10% of intact DETA NONOate remained in the microparticles after 1 month, indicating that the microparticles could not protect DETA NONOate from its decomposing under an ambient condition. 3.3.5. Differential Scanning Calorimetry (DSC) Study on PLGA Microparticles

1. Use differential scanning calorimetry (DSC) analysis to assess thermal properties of the compounds (see Note 12). 2. Examine samples, such as DETA NONOate (as raw powder), PLGA 5050, a mixture of DETA NONOate and polymer, and DETA NONOate-containing microparticle (w/o/wa ), using the Q100 DSC (TA Instruments, New Castle, DE, USA). 3. Place each sample (∼10 mg) in hermetically sealed aluminum pans and heated at a rate of 5◦ C/min up to 200◦ C under dynamic nitrogen (N2 ) atmosphere. The results of the DSC study showed that the microparticles loaded with DETA NONOate did not display any peaks but Tg at the temperature range of 50–150◦ C, indicating that DETA NONOate was not crystallized in the microparticles during the fabrication process and present in an amorphous from.

3.3.6. In Vitro NO Release Study

1. Make 100 mM stock solution citrate buffer as follows: 1.92 g citric acid in 100 mL ddH2 O. 2. Make 100 mM NaCitrate dehydrate stock buffer as follows: 14.7 g NaCitrate dihydrate to 500 mL ddH2 O. 3. Make a working solution of citric solution by combining 9 mL of 100 mM citric acid with 41 mL of 100 mM NaCitrate dehydrate. Bring to a final volume of 500 mL with ddH2 O. 4. Suspend the microparticles (5 mg) in 1 mL of citrate buffer solution (pH 4.0 or 5.5) or PBS (1×, pH 7.4). 5. Incubate the suspended microparticles at 37◦ C in the orbital shaker at a speed of 120 rpm. 6. At a predetermined time interval, take the samples and centrifuge at 12,000×g for 10 min. 7. Analyze the intact DETA NONOate in the supernatant by spectrophotometry at a wavelength of 252 nm.

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8. Digest the pellet using 1 M NaOH and measure the remaining amount of DETA NONOate in the microparticles. 9. Prepare all samples in triplicate and calculate the amount (%) of NO released from the microparticles by the following equation. NO released (%) = 100 − (% remaining DETA NONOate+ % DETA NONOate in the supernatant) The release profiles of NO from the microparticles were dependent on the fabrication methods and polymer compositions. Most of NO loaded in PLGA microparticles prepared by w/o/w method was released within the first 2 h, while the release rates of NO from those prepared by w/o/wa and w/o/o methods were further sustained over 6 h. Microparticles prepared by w/o/o displayed the initial burst release of NO with the amount of 30–50%, whereas those prepared by w/o/wa displayed that of 25–35% in 30 min. 3.4. Evaluation of the Film 3.4.1. Characterization of the GSNO Film According to the Drying Methods

3.4.2. Measurement of the GSNO Film Thickness

1. Prepare polymeric vaginal films containing GSNO by the solvent evaporation method (see Section 3.2) 2. Dry the polymer solutions containing GSNO under the conditions of either (i) a room temperature (25◦ C), (ii) the conventional heat drying methods (40 and 60◦ C), or (iii) a reduced pressure (20 mmHg) as well as a low temperature (15◦ C). The conventional heat drying methods (40 and 60◦ C) produced the film with a cloudy and tiny bubbled surface as well as about 70% loss of GSNO, indicating that GSNO quickly decomposed and released NO in a form of bubbles during the dry process (Table 13.2). The drying method under the condition of room temperature (25◦ C) enhanced the stability of GSNO in the film and loading efficiency up to 48% (see Note 13). The drying method under the condition of reduced pressure (20 mmHg) along with low temperature (15◦ C) in a vacuum (VirTis, Gardiner, NY, USA) produced the homogenous films with a reddish transparent surface and about a 71% loading efficiency (see Note 14). 1. Measure the thickness of each film at five different locations (center and four corners) using a micrometer screw gauge (Fowler Co., Japan). The mean value of five locations was used as a film thickness. The thickness of the film increased (119–194 μm) as the loading dose of GSNO decreased (30 to 5%), indicating that the thickness

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Table 13.2 The GSNO films prepared by various drying methods (cited from reference 8) Drying methodsa

Drying time (h)

Loading efficiency (%)

Physical characteristics

At 25◦ C

36

48 ± 2

Cloudy surface, phase separation

At 40◦ C At 60◦ C

12

29 ± 2

Cloudy pink surface with bubbles

8

30 ± 3

Cloudy pink surface with bubbles

Under a reduced pressureb

15

71 ± 2

Reddish, transparent, homogeneous surface

a The loading dose was 20 wt.% for all films. b Under 20 mmHg at 15◦ C.

Table 13.3 Characterization of the GSNO films with different loading doses (cited from reference 8) GSNO loadinga

Thicknessb (μm)

Loading efficiency (%)

Loading capacity (%)

30 wt.%

119 ± 10

51 ± 3

15.2 ± 0.9

20 wt.%

132 ± 18

71 ± 2

14.2 ± 0.5

10 wt.%

156 ± 15

83.7 ± 0.9

8.4 ± 0.1

5 wt.%

194 ± 15

94.9 ± 1.1

4.7 ± 0.1

a All films were dried under a reduced pressure. b n = 5.

of the film depends on the amount of polymer rather than the GSNO in the film (Table 13.3). 3.4.3. Assessment of Mechanical Properties of the GSNO Film

1. Evaluate the stress strain of the films using an Instron 5848 microtester (Norwood, MA, USA) equipped with Bluehill software and a 50 N load cell. 2. Position films (4 × 0.5 cm2 ) between two tensile grips at a distance of 1 cm and assess their stress strain at a rate of 1%/s at room temperature. The blank film has 79% elongation with good mechanical properties. Although the incorporation of GSNO into the film reduced stress and strain properties of the film, the GSNO film still retained the proper mechanical properties (see Note 15).

3.4.4. Assessment of Thermal Properties of the GSNO Film

1. Use differential scanning calorimetry (DSC) analysis to examine the thermal properties and nature of NO dispersion in the film (16). 2. Examine the thermal properties of samples, such as the pure GSNO, a blank film, and the GSNO film (20 wt.%), by

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comparing their melting points using Q100 DSC (TA Instruments, New Castle, DE, USA) (see Note 16). 3. Each sample was placed in hermetically sealed aluminum pans and heat at a rate of 5◦ C/min up to 200◦ C under dynamic N2 atmosphere. The peak representing the melting point of GSNO disappeared in the film containing GSNO, indicating that GSNO is fully dissolved and molecularly dispersed in the polymer film. 3.4.5. Assessment of Loading Efficiency and Loading Capacity of the GSNO Film

1. Assess the loading efficiency (LE) and loading capacity (LC) of GSNO in the final products to determine formulation properties of the polymeric film. 2. Cut, weigh, and place film samples in cold water to prevent GSNO loaded in the film from further decomposition. 3. Disintegrate and disperse the films in the solution by vortexing. 4. Centrifuge at 20,000×g for 20 min. Analyze the GSNO amount in the clear supernatant by spectrophotometry at a wavelength of 336 nm. 5. Calculate LE and LC from at least three different batches using the following equations. LE (%) =

LC (%) =

The amount of GSNO in the film × 100 The amount of initially loaded GSNO

The amount of GSNO in the film × 100 The total amount of polymers and GSNO

LE and LC are dependent on the initial loading amount and drying methods. As the initial loading amount increased, the LE in the films decreased, whereas the LC of GSNO increased (Table 13.3). 3.4.6. Stability Study of GSNO in the GSNO Film

1. Determine the stability of GSNO in the films in a temperature-humidity chamber (Hotpack, Philadelphia, PA, USA) under the conditions of (i) 25◦ C with the relative humidity of 60%, (ii) a refrigerator at 4◦ C, or (iii) a freezer at –20◦ C. 2. Keep all film samples in a glass bottle covered with the aluminum foil to protect them from the light. 3. Analyze the remaining amount of GSNO in the film after predetermined time intervals using spectrophotometry at a wavelength of 252 nm. There were no significant changes in the integrity of GSNO in the films at –20◦ C for 30 days. Around 6.5% of GSNO decomposed at 4◦ C after 30 days. The decomposition rate of GSNO was

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faster at 25◦ C than those at lower temperatures. Around 40% of GSNO decomposed within the first 10 days and about a half of GSNO remained intact for up to 30 days. 3.4.7. In Vitro NO Release Study from the GSNO Film

1. Examine the release profiles of NO to determine the optimal loading dose in the film (see Note 17). 2. Cut and immerse films in citrate buffer solution (pH 4.0 or 5.5) or PBS (pH 7.4) at 37◦ C. 3. Determine the remaining GSNO amount at a predetermined time interval as previously described. The amount of NO released from the films at time t ([NO]t ) was calculated by Equation [1]: [NO]t = [GSNO]0 − [GSNO]t

[1]

where [GSNO]0 is the initial concentration of GSNO and [GSNO]t is the concentration of GSNO at time t. 4. Calculate the amount of NO released from the film, from the amount of GSNO decomposed. Determine the NO released from the film using the first-order rate constant (k) obtained from the kinetic curve of the GSNO decomposition profile: [GSNO]t = [GSNO]0 Exp(−kt)

[2]

It was found that the release rates of NO from the GSNO films were slow for the first 1 h and afterward the NO release profiles from the films displayed the first-order kinetic. About 50% of NO were released within 10 h from the films loaded with 20% and 30 wt.% of GSNO, whereas less than 20% of NO were released from the films loaded with 5% and 10 wt.% of GSNO. 3.5. Cytotoxicity Study of NO Formulations

1. Perform cell viability studies to evaluate whether DETA NONOate or NO-releasing microparticles/films induce any cytotoxic effects on target cells (17). 2. Harvest VK2E6E7, vaginal epithelial cells, at 70% confluence (see Note 18). 3. Seed 2 × 103 cells/well were seeded in a 96 well plate pre-loaded with keratinized serum-free medium containing supplements. 4. Incubate the cells at 37◦ C under 5% CO2 and 95% humidity. 5. Replace the medium in the wells after 24 h with the experimental medium containing various amounts of DETA NONOate (0–5 mM) or the NO-releasing microparticles (equivalent to 0–5 mM of DETA NONOate). 6. Incubate for 6 h after which the medium in the wells was replaced with 100 μL of medium containing 20 μL of the

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Cell Titer 96 Aqueous assay solution. Incubate the plate at 37◦ C for 2 h. 7. Use drug-free medium as a control and load well with the medium only as a background control. 8. Measure the absorbance of the formazan products using a Beckman Coulter DTX 880 Multimode Detector (Beckman Coulter, Fullerton, CA, USA) at 485 nm. Neither DETA NONOate (0.1–2.5 mM) nor NO-releasing microparticles (equivalent to 0.1–2.5 mM of DETA NONOate) caused any cytotoxicities to vaginal cells with 6 h of treatment. 3.6. In Vivo Study: Measurement of Changes in the Vaginal Blood Flow by NO Formulations

1. Use female Wistar rats (Charles River, Canada) weighing between 250 and 270 g for physiological and pathological studies for vaginal or cardiovascular blood flow models following exposure to NO-releasing microparticles or films (9, 18) (see Note 19). 2. Administer Ketamine hydrochloride (35 mg/kg) and xylazine (3.5 mg/kg) by i.m. to maintain anesthesia throughout the experiment. 3. Use Laser Doppler Flowmetry (PeriFlux 4001 Master, Perimed, Stockholm, Sweden) equipped with a PF 415:1 probe (tip diameter, 0.7 mm) to measure vaginal blood flow. 4. Place the Laser Doppler probe into vaginal lumen facing the anterolateral vaginal wall 15–17 mm distal from the vulva as described previously (9). 5. Calibrate the Laser Doppler flowmetry with external standards using the Periflux 1000 Calibration Device prior to the experiment (see Note 20). 6. Dissolve non-formulated DETA NONOate or microparticles in a normal saline solution immediately before use. 7. Insert a polyethylene tube (Intramedic PE50 or PE190) along with the probe into the vaginal vault to facilitate the injection of fluid. 8. Euthanize with an i.m. injection of pentobarbital (100 mg/kg). Non-formulated DETA NONOate (2 mg) significantly enhanced the vaginal blood flow for only 10 min after application, whereas NO-releasing microparticles containing 2 mg of DETA NONOate significantly enhanced the vaginal blood flow at 5 min after its administration and maintained the enhanced vaginal blood flow up to 120 min (p < 0.05 versus baseline, n = 3). The maximum percentage change in vaginal blood flow of 163 ± 13 was obtained at 30 min after application (see Note 21).

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4. Notes 1. 5050 and 7525 means the ratio of lactides and glycolides. I.V. stands for Inherent Viscosity. The degradation rate of the foam in LPGA can be controlled by careful selection of the copolymer composition and molecular weight. Microparticles made of PLGA 7525 showed a slower release profile of NO than those of PLGA 5050. 2. During the initial screening process, we found that diethylenetriamine diazeniumdiolate (DETA NONOate) is suitable for PLGA polymers (i.e., microparticle formulations) and S-nitrosoglutathione (GSNO) for polymer mixtures (i.e., films). 3. The reduced form of glutathione used was based on the fact that highly reduced forms of glutathione are strong antioxidants. 4. The size and entrapment efficiency of microparticles displayed a wide variance according to the fabrication methods. The oil/water (o/w) technique seemed to be superior to the oil/oil (o/o) approach and the solvent evaporation method was found to be more efficient than the solvent extraction method due to the higher encapsulation rate. Microparticles made by double emulsion methods (w/o/w, w/o/wa and w/o/o) had greater particle sizes and entrapment efficiency than those made by single emulsion methods (o/w and o/o). 5. Three doses (w/w) of DETA NONOate were used to evaluate the effects of loading dose on NO release profiles. 6. The decomposition rate of DETA NONOate in the microparticles at 25◦ C could be accelerated by the acidic nature of PLGA which consists of acidic degradation components (lactic acid and glycolic acid). Since the NO donor is less stable in acidic conditions, alkaline water (wa ) used in the product of (w/o/wa ) enhanced the stability of the NO donor in microparticles and subsequently prolonged the drug release period. 7. This process is integral to maintain biocompatibility of the microparticles. 8. The removal of the air bubbles entrapped is essential to produce a smooth and homogeneous surface morphology. 9. Careful peeling is needed to avoid any accidental break or surface damage to the film.

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10. AFM characterizes the microparticle size more accurately than SEM when the tip curvature radius is much smaller than the microparticle size. 11. The half-life of DETA NONOate at a physiological pH is short, denoting 22 and 56 h at 37 and 25◦ C, respectively. 12. DSC is used to determine whether or not the NO donor is homogeneously distributed and thermal properties of the microparticles are maintained. PLGA had a glass transition temperature (Tg ) at around 46◦ C and DETA NONOate had an endothermic melting peak at 104◦ C. The physical mixture of polymer and drug exhibited both Tg and melting point. 13. It was occasionally observed that the phase separation of GSNO occurred at a cloudy surface of the polymeric film prepared at room temperature. 14. GSNO films prepared under the reduced pressure (20 mmHg) as well as a low temperature (15◦ C) were selected and used for subsequent experiments. 15. It was reported that a range of 20–100% of elongation of the film was considered flexible but hard enough to endure mechanical stress in nature (24). The lowered stress and strain values at the break of the GSNO film are mainly due to the presence of the small amount of polymers in the film as the polymer network defines the mechanical nature of the film. 16. GSNO displayed a strong endothermic peak (i.e., a melting point) at 195◦ C. A broad endothermic peak was observed below 100◦ C in both blank film and GSNO film, which can be attributed to the evaporation of moisture absorbed in the films. 17. The NO release profile from the films reflects the spontaneous decomposition of GSNO both inside and outside of the films. 18. K2E6E7, whose physiological properties are very similar to those of human vagina cells, has been frequently used for evaluation of drug permeation profiles and uptake mechanisms. 19. The rat model has been widely used for this purpose due to ease of handling and sensitivity in blood flow changes. 20. The maximum change in vaginal blood flow was calculated and compared with baseline value (i.e., the value at 0 min) induced by normal saline solution (i.e., the control). 21. The results support that NO-releasing systems could improve blood flow and can be used for the treatment of cardiovascular diseases and female sexual arousal disorder.

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Acknowledgment This work was partly supported with a grant from the Missouri Life Science Research Board (09-1117). The author would like to thank Dr. Jin-Wook Yoo for his contribution to the formulation development and evaluation processes and Dr. Peter Smith in KUMC for lending us the laser Doppler. References 1. Naseem, K. M. (2005) The role of nitric oxide in cardiovascular diseases. Mol Aspects Med 26, 33–65. 2. Traub, O., Vanbibber, R. (1995) Role of nitric-oxide in insulin-dependent diabetesmellitus related vascular complications. West J Med 162, 439–445. 3. Uckert, S., Ehlers, V., Nuser, V., Oelke, M., Kauffels, W., et al. (2005) In vitro functional responses of isolated human vaginal tissue to selective phosphodiesterase inhibitors. World J Urol 23, 398–404. 4. Kim, S. W., Jeong, S. J., Munarriz, R., Kim, N. N., Goldstein, I., Traish, A. M. (2003) Role of the nitric oxide-cyclic GMP pathway in regulation of vaginal blood flow. Int J Impot Res 15, 355–361. 5. Ignarro, L. (2000) Nitric Oxide: Biology and Pathobiology. Academic, San Diego, IL. 6. Ferrara, D., Zaslau, S. (2007) Success of sildenafil treatment in neurogenic female sexual dysfunction caused by L5-S1 intervertebral disk rupture: a case report. Int J Urol 14, 566–567. 7. Yoo, J. W., Lee, J. S., Lee, C. H., Lee, C. H. (2010) Characterization of nitric oxidereleasing microparticles for mucosal delivery. J Biomed Mat Res 92, 1233–1243. 8. Yoo, J. W., Acharya, G., Lee, C. H. (2009) In vivo evaluation of vaginal films for mucosal delivery of nitric oxide. Biomaterials 30, 3978–3985. 9. Yoo, J. W., Choe, E. S., Ahn, S. M., Lee, C. H. (2010) Pharmacological activity and protein phosphorylation caused by nitric oxide-releasing microparticles. Biomaterial 31, 552–558. 10. Hrabie, J., Klose, J., Wink, D., Keefer, L. (1993) New nitric oxide-releasing zwitterions derived from polyamines. J Org Chem 58, 1472–1476.

11. Pacher, P., Mabley, J. G., Liaudet, L., Evgenov, O. V., Southan, G. J., Abdelkarim, G. E., et al. (2003) Topical administration of a novel nitric oxide donor, linear polyethylenimine-nitric oxide/nucleophile adduct (DS1), selectively increases vaginal blood flow in anesthetized rats. Int J Impot Res 15, 461–464. 12. Bilati, U., Allemann, E., Doelker, E. (2005) Poly(D,L-lactide-co-glycolide) proteinloaded nanoparticles prepared by the double emulsion method – processing and formulation issues for enhanced entrapment efficiency. J Microencapsul 22, 205–214. 13. Dubernet, C. (1995) Thermoanalysis of microspheres. Thermochim Acta 248, 259–269. 14. Arcoleo, V., Liveri, V. T. (1996) AFM investigation of gold nanoparticles synthesized in water/AOT/n-heptane microemulsions. Chem Phy Let 258, 223–227. 15. Pauwels, R., Balzarini, J., Baba, M., Snoeck, R., Schols, D., Herdewijn, P., et al. (1988) Rapid and automated tetrazolium-based colorimetric assay for the detection of antiHIV compounds. J Virol Methods 20, 309–321. 16. Dubernet, C. (1995) Thermoanalysis of microspheres. Thermochim Acta 248, 259–269. 17. Berridge, M. V., Tan, A. S. (1993) Characterization of the cellular reduction of 3-(4,5-dimethylthiazol-2-yl)-2,5diphenyltetrazolium bromide (MTT): subcellular localization, substrate dependence, and involvement of mitochondrial electron transport in MTT reduction. Arch Biochem Biophys 303, 474–482. 18. Treisman, R. (1996) Regulation of transcription by MAP kinase cascades. Curr Opin Cell Biol 8, 205–215.

Chapter 14 Nitric Oxide Releasing Nanoparticle Synthesis and Characterization George Han, Adam J. Friedman, and Joel M. Friedman Abstract While the potential applications of nitric oxide for both understanding human physiology and treating disease are far reaching, the development of a reliable, cost-effective, and practical sustained delivery system for nitric oxide has yet to emerge. Using a sol–gel/glass hybrid system, we have demonstrated controlled, sustained release of nitric oxide from a stable, dry powder. Upon exposure to an aqueous environment, the material begins releasing therapeutic levels of nitric oxide over several hours to days, making it an ideal material for evaluation of nitric oxide efficacy for both clinical and research applications. Key words: Nitric oxide, sol–gel, nanoparticles, chitosan, sustained release.

1. Introduction The role of nitric oxide (NO) in physiology has been extensively studied in recent decades, since the discovery of its role in vasodilation. It has become apparent that nearly every physiological system is linked to NO in some way, with vascular biology (1), neuroscience (2), and immunology (3) representing only a few of the systems for which NO is an important regulatory factor. NO’s vast and diverse effects are dependent on localization and duration of production. For example, as a signaling molecule, NO is not only the primary regulator of vascular tone throughout the body but also involved as a neurotransmitter in the cerebral cortex and cerebellum where it has been implicated in processes including longterm potentiation of memories (4). Even within the brain itself, NO has seemingly opposing effects, mediating differentiation and regeneration of neurons (5, 6) while also serving as a neurotoxic agent as seen in Huntington’s and Alzheimer’s diseases (7, 8). H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_14, © Springer Science+Business Media, LLC 2011

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With such widespread effects intimately linked to finely regulated control of NO levels, it is not surprising that progress translating NO into a clinical therapy has been slow and difficult. In order to address the challenges in administering NO as a therapeutic, we have developed a hybrid nanoparticle system combining features of both silane-based hydrogels and sugar-derived glassy matrices. The result is a material that exhibits hydrationdependent porosity while remaining highly robust structurally, thus allowing for slow diffusion of NO out of the material over time (9). The composition of the nanoparticles allows for retention of NO within the dry particles, as well as provides for slow sustained release of therapeutic levels of NO over long time periods when exposed to moisture/water. Unlike many of the current NO releasing materials, NO release from the nanoparticles does not require either chemical decomposition or enzymatic catalysis, only exposure of the particles to water. The release profile for NO is easily tuned through straightforward manipulation of the relative concentrations of the components used in preparing the hydrogel/glass composite that are the basis for the nanoparticle platform. To date, several translational applications have been demonstrated. Topical application of the NO releasing nanoparticles to both uninfected and infected wounds with drug-resistant Staphylococcus aureus demonstrated accelerated wound closure and clearance of bacterial invasion (10). Modulation of vasoactivity has also been investigated as a target for NO nanoparticle therapy in order to combat diseases relating to endothelial dysfunction. Using erectile dysfunction as a model for endothelial pathology, we demonstrated that NO nanoparticles increased erectile function when applied to the penis of a rat model (11). Furthermore, topical application to the chest wall of an animal hypertension model resulted in a sustained decrease in mean arterial pressure with minimal systemic adverse effects. Numerous applications are currently being investigated and are seemingly limited only by the myriad applications of NO itself. The potential is clear – NO nanoparticles can function as a therapeutic agent for inflammatory, infectious, vascular/cardiovascular, and thrombotic disorders, as well as a very promising tool to promote our understanding of NO signaling mechanisms.

2. Materials 2.1. Synthesis of NO-Releasing Nanoparticles

1. Tetramethyl Orthosilicate (TMOS). 2. Solution of HCl between 0.2 and 0.5 mM, depending on the batch of TMOS used (see Note 1).

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3. Phosphate buffer (0.05 M) 7.4 pH. 4. Sodium nitrite is dissolved in the phosphate buffer at 0.9 M or less, if a lower total amount of NO is desired (see Note 2). 5. Glucose is added to the 40 mg/mL 0.05 M phosphate buffer 7.4 pH. 6. Polyethylene glycol (PEG) available in average molecular weights of 200–6,000 and selected based on rate of release of NO desired (see Note 3). 7. 5 mg/mL chitosan is dissolved in 1% acetic acid to form a stock solution of chitosan. 8. Sonicator to hydrolyze TMOS. 9. Flask lyophilizer (Virtis benchtop) to dry the solid gel. Other brands/types may be used as well. 10. Ceramic mortar and pestle. 11. Planetary ball mill (Fritsch Pulverisette). 2.2. NO Release Measurement

1. Nitric oxide (gas tank). 2. Gastight syringe (Hamilton company, Reno, NV). 3. Apollo 4000 Free Radical Detector (World Precision Instruments [WPI], Sarasota, FL). This is a four-channel amperometric NO detector, although they also produce a onechannel version that can be used as well. 4. ISO-NOP Sensor (WPI). This sensor is their original NO sensor and although newer variants have been introduced, this one remains the most robust and is the most selective toward NO as opposed to other compounds such as nitrite and it is therefore the most appropriate for measuring NO release from the nanoparticles. 5. Phosphate buffer (0.05 M) 7.4 pH.

2.3. NO-Releasing Nanoparticle Administration

1. Normal saline. 2. Carboxymethylcellulose (Hercules corp., Wilmington, DE).

3. Methods The synthesis of a NO releasing material from a nitrite precursor depends on successful reduction of nitrite to NO, while retaining the latter in the material. Since NO is a volatile gas that reacts with atmospheric gases quickly, storage of NO should be in a form where it is not able to interact with atmospheric gases.

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Traditionally, this would call for careful storage protocols and synthesis procedures. However, using the characteristic of a sol–gel to entrap and sequester compounds and the ability of glasses to form robust materials, we have developed a system to both form NO from nitrite and store it over extended time periods. The only requirement is that the stored material be kept away from moisture, in a sealed container. The use of silane precursors for encapsulating molecules and even cells has been well studied, allowing for diffusion of small molecules through pores in the hydrogel. We sought to add other characteristics to form a more robust material, adding PEG to plug the pores and using chitosan to endow glass-like properties to the material. Additionally, the ability of chitosan to facilitate electron transfer, open gap junctions, and kill pathogens itself has proven to be beneficial. Chitosan is also advantageous in terms of forming NO, penetrating through the skin and providing an antibacterial action. Glucose is used as an electron donor to facilitate reduction of nitrite to NO, although we have recently demonstrated that its inclusion is not necessary for efficient reduction of nitrite to NO. In this sense, the chitosan itself may be facilitating reduction of nitrite to NO, in addition to other mechanisms of reduction. Measurement of NO is also somewhat complicated due to its promiscuous interactions with other compounds owing to its free radical nature. Previously, Griess reagent has been used as a measure of NO by first converting it to nitrite, but this is not a precise estimate and is sensitive to nitrite as well. While we have directly tested our material with the Griess assay and saw no signal, the minimum concentration of nitrite that the Griess assay is able to measure is 1 μM, revealing a lack of sensitivity with the Griess assay. Diaminofluorescein (DAF) has also been used to measure NO concentration (12) and while we have used this technique in our laboratory, we found that it is relatively complicated and inconsistent. This is due to the reaction with N2 O3 , which requires the participation of atmospheric oxygen, confounding the direct measurement of NO. Furthermore, over longer time periods, saturation and photobleaching may become an issue when using a fluorescent reagent and therefore its use in measuring NO output from the nanoparticle is not recommended. We could obtain excellent results from the Apollo 4000 NO Detector from WPI, due to its ability to monitor real-time NO release at very low concentrations. Additionally, it can directly measure NO in a wide variety of solutions. However, it is sensitive to light, pH, and temperature, so these variables must be monitored during use. WPI makes various probe tips for NO sensing and certain probes are more suited to specific applications. The ISO-NOP sensor, while one of the largest and the oldest design, has excellent selectivity toward NO. While other tips may be more

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ideal for conditions such as measurement in cell culture, where flexible tips down to a few microns in diameter are available, the ISO-NOP is well-suited for measurement in solution. 3.1. Synthesis of NO-Releasing Nanoparticles

1. The TMOS should be sonicated for hydrolysis to occur. For every 5 mL TMOS, add 560 μL 0.2–0.5 mM HCl (see Note 1) and 605 μL distilled H2 O (see Note 4). Sonicate the TMOS in a small glass bottle with a stopper or tightly sealed with parafilm (see Note 5) for 1 h or until the solution is fully hydrolyzed and homogenous (see Note 6). 2. Prepare the nitrite/glucose solution in phosphate buffer near the time when the hydrolysis will be done. For every 5 mL of stock TMOS (about 6 mL after hydrolysis, including water and acid volume), 60 mL of buffer will be needed. Add 0.9 M sodium nitrite (see Note 2) to the buffer solution while stirring and when fully mixed, add the glucose. Set aside when all components have dissolved. 3. Once the hydrolysis of TMOS is finished, first add PEG to the solution and then chitosan. PEG (see Note 3) is added at a ratio of 1 mL per 20 mL of buffer. Chitosan (see Note 7) is also added at a ratio of 1 mL per 20 mL of buffer. Lastly, add the hydrolyzed TMOS at 1 mL per 10 mL of buffer. Cover loosely with parafilm and allow gelation to occur, which may take minutes to hours. 4. After waiting at least several hours or overnight and ensuring that the gel has formed fully, press the solid gel between layers of paper towels to remove excess methanol. Place the resultant material in a beaker, flask, or tube and lyophilize for at least 24 h or until fully dried (see Note 8). 5. Take the dry powder material and roughly grind with a mortar and pestle both to make the powder finer and to make sure that there are no areas of moisture remaining in the sample. Place the sample in the well of the ball milling apparatus and cover. Ball mill for 60 min at 3.5 g (see Note 9). Remove sample from well and place into sealable container, making sure to seal tightly to avoid contact with atmospheric moisture. A sample TEM image of the final product is shown in Fig. 14.1.

3.2. NO Release Measurement

1. Make saturated NO solution by first purging an aliquot of water with Nitrogen or Argon gas, then bubbling in NO gas for at least 20 min. Run the gas through a solution of NaOH (2–5 M) first to remove higher oxides. This solution will have a concentration of NO at 1.9 mM. Alternatively, a NO donor such as S-Nitroso-N-acetylpenicillamine may be used to calibrate the sensor (see WPI documentation for instructions).

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Fig. 14.1. NO-releasing nanoparticles as viewed with TEM. The scale bar represents 100 nm.

2. Plug in sensor to Apollo 4000 instrument and submerge probe tip in phosphate buffer, while stirring. Calibrate the sensor by adding known volumes of saturated NO solution with a gastight syringe and calculating what the concentration of NO is with each addition. Each addition of saturated NO solution should be initiated when the signal has reached a plateau (see WPI documentation for detailed instructions). 3. Remove sensor from calibration solution, wash, and place in a beaker with fresh buffer. After the signal has reached a stable baseline (see Note 10), add a known weight of NO-releasing nanoparticles (we use 100 mg per 20 mL buffer). Record data. 4. Compare to calibration curve to convert picoampere readings from Apollo 4000 to NO concentrations. Final sample data readout is shown in Fig. 14.2. 3.3. NO-Releasing Nanoparticle Administration

1. NO-releasing nanoparticles (NO-nps) have broad applications, both in vitro and in vivo. When testing in vitro for cell culture, the powder may be added directly to cell culture plates (see Note 11). 2. For topical in vivo application, carboxymethylcellulose (CMC) may be used as a bulking agent. A 1–2.5% solution of CMC is ideal for use on a topical wound, as it keeps the

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Fig. 14.2. Nitric oxide release from nanoparticles. Data were collected for over 1 day, revealing sustained release of NO during this period.

application localized. Just prior to administration, add the desired weight of NO-nps to a solution of CMC and then apply. 3. For IV or IP administration, add NO-nps to saline and then vortex and/or sonicate the solution briefly. The sample should be administered with the largest bore needle appropriate for the application, to prevent clumping of the material at the needle tip.

4. Notes 1. The concentration of acid necessary to achieve hydrolysis depends on the batch of TMOS used. Currently, SigmaAldrich has a few batches released every year, with each batch showing slightly different characteristics, mostly concerned with optical transparency, which is not an issue with NO-np synthesis. 2. Sodium nitrite is the precursor to NO, so if a reduced final concentration of NO is desired, reduce the amount of sodium nitrite initially used. Above 0.9 M, the process of reduction and retention becomes less efficient, therefore using concentrations above 0.9 M is not recommended. 3. As PEG is used to affect the pore size in the sol–gel network, it is the primary effector of release rate. Higher

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molecular weight PEG’s will cause the pores around it to be similarly larger, and thus NO release is truncated at a higher concentration over a shorter time period. We find that using PEG 400–600 MW offers a good balance between release rate and duration of release. 4. All water used should be deionized at a resistivity of 18 M cm. Care should be taken to test for copper contamination, as it is widespread and can affect the balance of NO product formed. 5. If using parafilm, wrap the bottle top with parafilm many times to ensure a tight seal. This method may be preferred to rubber stoppers as material may condense on the rubber and become problematic. 6. Full hydrolysis depends on many variable factors, including the batch of TMOS, the acid used, and time. The completely hydrolyzed product should be clear and with brief, light shaking should appear homogenous. 7. Chitosan should be added immediately before the TMOS, as its slightly acidic nature may initiate some NO formation. This will be lost prior to gelation initiated by the TMOS. The pH of the chitosan solution should be no lower than 5.5; otherwise, less acetic acid should be used (as little as possible to fully dissolve the chitosan). We have found that the practical grade chitosan available from Sigma-Aldrich is the easiest to work with and readily dissolves at low concentrations of acetic acid. Furthermore, fresh chitosan batches should be made every 1–2 months, as it is known to break down and degrade over time in solution. 8. The lyophilizer used should have enough vacuum to pull all of the moisture out of the towel-dried gels within 48 h. The presence of other flasks on the lyophilizer may affect performance. 9. The Planetary Ball Mill is available with a wide range of wells and balls to achieve grinding. Ideally, the smallest well that accommodates the amount of sample being ground should be used. Furthermore, balls made from synthetic materials (not metal) are preferred since they generate less heat during the milling process. 10. Many factors can make generating a steady baseline on the Apollo 4000 challenging. Among them are light, pH, and temperature, as mentioned previously. Of these, temperature is important because slight variations in laboratory temperature, as common in day–night cycles, can generate a drift in the signal that may be confused with a genuine NO signal. The Apollo 4000 instrumentation supports attachment of a separate temperature probe to follow

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temperature at the measurement site, which can be correlated to the signal. Additionally, an isolated electrical source will help to create a stable signal. Plugging the machine on the same power strip as a stirrer/hot plate, for example, is not recommended. The equipment may be enclosed in a glove box purged with nitrogen to exclude loss of signal due to NO reaction with atmospheric gases. 11. Different cell types have different sensitivity to NO-nps. For example, for human fibroblasts, do not exceed 50 mg of NO-nps for a standard 10 cm plate. For primary neuron cultures, do not add more than 5 mg of NO-nps. References 1. Palmer, R. M. J., Ferrige, A. G., Moncada, S. (1987) Nitric oxide release accounts for the biological activity of endotheliumderived relaxing factor. Nature 327, 524–526. 2. Murad, F. (1998) Nitric oxide signaling: would you believe that a simple free radical could be a second messenger, autacoid, paracrine substance, neurotransmitter, and hormone? Recent Prog Horm Res 53, 43–59. 3. Bogdan, C. (2001) Nitric oxide and the immune response. Nat Immunol 2, 907–916. 4. Malen, P. L., Chapman, P. F. (1997) Nitric oxide facilitates long-term potentiation, but not long-term depression. J Neurosci 17, 2645–2651. 5. Northington, B. F. J., Tobin, J. R., Harris, A. P., Traystman, R. J., Koehler, R. C. (1997) Developmental and regional differences in nitric oxide synthase activity and blood flow in the sheep brain. J Cereb Blood Flow Metab 17, 109–115. 6. Verge, V. M. K., Xu, Z., Xu, X. J., Wiesenfeld-Hallin, Z., Hokfelt, T. (1992) Marked increase in nitric oxide synthase mRNA in rat dorsal root ganglia after peripheral axotomy: in situ hybridization and functional studies. PNAS 89, 11617–11621. 7. Browne, S. E., Ferrante, R. J., Beal, M. F. (1999) Oxidative stress in Huntington’s disease. Brain Pathol 9, 147–163.

8. Vodovotz, Y., Lucia, M. S., Flanders, K. C., Chesler, L., Xie, Q. W., Smith, T. W., et al. 1(1996) Inducible nitric oxide synthase in tangle-bearing neurons of patients with Alzheimer’s disease. J Exp Med 184, 1425–1433. 9. Friedman, A. J., Han, G., Navati, M. S., Chacko, M., Gunther, L., Alfieri, A., Friedman, J. M. (2008) Sustained release nitric oxide releasing nanoparticles: characterization of a novel delivery platform based on nitrite containing hydrogel/glass composites. Nitric Oxide 19, 12–20. 10. Martinez, L. R., Han, G., Chacko, M., Mihu, M. R., Jacobson, M., et al. (2009) Antimicrobial and healing efficacy of sustained release nitric oxide nanoparticles against Staphylococcus aureus skin infection. J Invest Dermatol 129, 2463–2469. 11. Han, G., Tar, M., Kuppam, D. S., Friedman, A., Melman, A., Friedman, J., et al. (2010) Nanoparticles as a novel delivery vehicle for therapeutics targeting erectile dysfunction. J Sex Med 7, 224–233. 12. Kojima, H., Nakatsubo, N., Kikuchi, K., Kawaraha, S., Kirino, Y., Nagoshi, H., et al. (1998) Detection and imaging of nitric oxide with novel fluorescent indicators: diaminofluoresceins. Anal Chem 70, 2446–2453.

Chapter 15 NOS Antagonism Using Viral Vectors as an Experimental Strategy: Implications for In Vivo Studies of Cardiovascular Control and Peripheral Neuropathies Beihui Liu, James Hewinson, Haibo Xu, Francisco Montero, Carmen R. Sunico, Federico Portillo, Julian F.R. Paton, Bernardo Moreno-López, and Sergey Kasparov Abstract Nitric oxide, a free gaseous signalling molecule, has attracted the attention of numerous biologists and has been implicated in the regulation of the cardiovascular, nervous and immune system. However, the cellular mechanisms mediating nitric oxide modulation remain unclear. Upregulation by gene overexpression or down-regulation by gene inactivation of nitric oxide synthase has generated quantitative changes in abundance thereby permitting functional insights. We have tested and proved that genetic nitric oxide synthase antagonism using viral vectors, particularly with dominant negative mutants and microRNA 30-based short hairpin RNA, is an efficient and effective experimental approach to manipulate nitric oxide synthase expression both in vitro and in vivo. Key words: Nitric oxide (NO), nitric oxide synthase (NOS), viral vectors, dominant negative mutants, RNA interference, cardiovascular control, nucleus tractus solitarii (NTS), peripheral neuropathy.

1. Introduction Currently, the most popular approach to investigate gene function is to generate a knock-out animal (typically a mouse) in which the resulting phenotype might give clues regarding gene function. However, this approach is costly, time-consuming and in some cases the function of targeted genes might not be determined because of embryonic lethality or maturational H.O. McCarthy, J.A. Coulter (eds.), Nitric Oxide, Methods in Molecular Biology 704, DOI 10.1007/978-1-61737-964-2_15, © Springer Science+Business Media, LLC 2011

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compensation (1, 2). These drawbacks limit its value in physiological genomics especially when it comes to understand highly complicated central nervous system pathways. In neuroscience experimentation, it is often essential to restrict the genetic intervention to specific cell types within a certain nucleus and to have temporal control over the expression of the transgenes. One way to meet all these requirements is to use somatic gene transfer targeted towards the selected nuclei in order to increase or decrease expression of a particular gene. However, transfer of genes into an intact brain in vivo remains a challenging task. While transfection of cell lines with plasmids is an efficient process particularly through the use of various chemical reagents to help permeability, transduction of mature neurones in a living brain is either resistant to these procedures or become damaged. However, viral vectors can effectively deliver genes into brain cells and even integrate them into the host genome for long-term expression making this a highly attractive approach (3, 4). Moreover, viral vectors have also been used successfully for the delivery of new genomic tools such as small interference RNAs both in vitro and in vivo (5, 6). In this chapter, we illustrate the application of viral tools for studies of nitric oxide (NO) function in peripheral nerve injury and central cardiovascular control (specifically at the level of NTS). NO function is dependent on dynamic regulation of its biosynthetic enzyme, NOS. There are three types of NOS, neuronal nitric oxide synthase (nNOS), endothelial nitric oxide synthase (eNOS) and inducible nitric oxide synthase (iNOS) (7). Of the three NOS isoforms, nNOS and eNOS constitute the predominant source of NO in the rat brain. Therefore, the targets of interest were either nNOS or eNOS. 1.1. Molecular Tools for eNOS and nNOS Antagonism

There are several strategies to achieve a full or partial “loss of function” using viral transgenesis. Here, we illustrate two – the expression of a dominant negative protein and a microRNA-based (miRNA) short hairpin RNA (shRNA) expression (Fig. 15.1).

1.1.1. Dominant Negative Approach

A dominant negative protein must in some way interfere with the target protein or its function. To suppress eNOS activity, a recombinant adenoviral vector (AVV) was used to express a truncated (T) form of eNOS (TeNOS) under the control of the human cytomegalovirus (CMV) promoter (8). TeNOS lacks catalytic activity but retains the NH2 -terminal sequences required for cotranslational NH2 -terminal glycine myristoylation (9) and membrane localization (10). Thus, TeNOS acts as a dominant negative inhibitor of wild-type eNOS activity through heterodimerization with the native protein (Fig. 15.1). As a viral control, AVV-CMVeGFP expressing enhanced green fluorescent protein (eGFP) was used (11, 12). Schematic representations of the two AVVs are

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Fig. 15.1. a: TeNOS acts as a dominant negative inhibitor of wild-type eNOS activity through heterodimerization with the native protein. b: A miR30-based shRNA expression vector. GFP and miR30-shRNA were co-transcribed by a polymerase II promoter and processed by Drosha/Dicer into functional reporter GFP and shRNA targeting the gene of interest. Mature shRNA is then loaded into RNA-induced silencing complex (RISC), which directs gene silencing through mRNA degradation or translational repression.

shown in Fig. 15.2a. Protocols for AVV preparation including the amplification and titration are described in Section 3.1. This vector has been used in two types of experiments – to assess the role of eNOS in central cardiovascular control at the level of NTS and to study the role of eNOS in peripheral neuropathy. Based on previous studies, we hypothesized that NO released from the endothelium in NTS increases inhibitory synaptic transmission affecting long-term blood pressure regulation (13). We injected AVV-CMV-TeNOS into the rat NTS and determined arterial pressure using radio-telemetry. We were able to demonstrate that a reduction in eNOS activity enhanced baroreceptor reflex function in normotensive rats and also lowered blood pressure in the spontaneously hypertensive rat (SHR) (14, 15) (Fig. 15.3b). These results were consistent with our hypothesis of vascular-neuronal signalling whereby ligands in the blood or produced within NTS activate eNOS located within the blood–brain barrier resulting in diffusion of NO to nearby neuronal networks (16). An important component in any in vivo gene transfer study is to demonstrate the cell type in which the transgene has been expressed. We previously demonstrated (14) that an antibody for endogenous eNOS also detected TeNOS. We showed upregulation of immunofluorescence for eNOS in animals that had received AVV-CMV-TeNOS relative to those receiving AVVCMV-eGFP. Other methods used to demonstrate upregulation

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Fig. 15.2. Schematic representation of viral vectors described in this chapter. a: Adenoviral vectors AVV-CMV-TeNOS and AVV-CMV-eGFP. ITR, adenoviral inverted terminal repeats; hCMV, human cytomegalovirus promoter; TeNOS, a truncated form of eNOS; eGFP, enhanced GFP reporter gene; b: Lentiviral vectors LVV-Tretight-GFP-miR30-shRNA/nNOS and LVVmCMV/SYN-tTA. LTR, lentiviral long terminal repeats; Tretight, a modified tetracycline and Dox-responsive promoter derived from pTRE-tight (Clontech); GFP, green fluorescence protein; miR30-shRNA/nNOS, miR30-based shRNA targeting rat neuronal nitric oxide synthase gene; SYN, human synapsin 1 promoter (470 bp); mCMV, minimal CMV core promoter (65 bp); GAL4BDp65, a chimaeric transactivator consisting of a part of the transactivation domain of the murine NF-κBp65 protein fused to the DNA-binding domain of GAL4 protein from yeast; WPRE, woodchuck hepatitis post-transcriptional regulatory element.

of transgene expression include western blotting or assays to measure NO production. Crushing of a motor nerve, such as the XIIth nerve, causes complete axotomy but unlike transection, it preserves the endoneural tube, providing neurotrophic support and a physical guide for the proximal axonal ends (17, 18). A few weeks after nerve crushing, muscle reinnervation takes place without significant loss of neurones. Therefore, traumatic injury by nerve crushing is a suitable model for the identification of molecules that regulate degenerative and regenerative processes. eNOS is overexpressed in vasa nervorum of the distal stump and around the injury site (19). Crushing the XIIth nerve induces an immediate and complete suppression of neuromuscular connectivity as demonstrated by the complete absence of the compound muscle action potential (CMAP) in the tongue after electrical stimulation of the injured nerve at the proximal portion. CMAP is completely absent 1, 3 or 7 days after crushing, even when supramaximal (>0.1 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 <0.25 g is acceptable between pairs of tubes. 37. Lentivirus pellets will not initially resuspend homogenously. Overnight incubation at 4◦ C aids the dissociation of virus particles. 38. Avoid introducing bubbles during the resuspension of the virus pellet. Limit freeze–thaw cycles as this will reduce virus titre significantly.

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39. The reaction solution is light sensitive, so keep tubes and plates wrapped in foil. 40. It may be difficult to differentiate between positively stained neighbouring cells infected with higher concentrations of virus; therefore, it is recommended that such dilutions of virus are not used in titre calculations. 41. Make sure that both sides and the bottom of the plates and spacers are flush, otherwise the gel will leak. 42. Ensure there are no bubbles in the separating gel. 43. Ensure the separating gel is smooth and even. 44. Ensure there are no bubbles in the stacking gel. 45. The buffer should not leak from the inner chamber. 46. Ensure the bottom of the gel is submerged in running buffer. 47. Roll a pippette on the filter paper to remove trapped air between the filter papers and PVDF membrane to ensure efficient transfer. 48. Cooling is essential during the transfer stage to prevent heat-induced damage to the apparatus and the experiment. 49. The exposure time may vary from 30 s to 3 min. References 1. Kasparov, S., Teschemacher, A. G., Hwang, D. Y., Kim, K. S., Lonergan, T., Paton, J. F. (2004) Viral vectors as tools for studies of central cardiovascular control. Prog Biophys Mol Biol 84, 251–277. 2. Aagaard, L., Rossi, J. J. (2007) RNAi therapeutics: principles, prospects and challenges. Adv Drug Deliv Rev 59, 75–86. 3. Thomas, C. E., Ehrhardt, A., Kay, M. A. (2003) Progress and problems with the use of viral vectors for gene therapy. Nat Rev Genet 4, 346–358. 4. Papale, A., Cerovic, M., Brambilla, R. (2009) Viral vector approaches to modify gene expression in the brain. J Neurosci Methods 15, 1–14. 5. Snove, O., Jr., Rossi, J. J. (2006) Expressing short hairpin RNAs in vivo. Nat Methods 3, 689–695. 6. Paddison, P. J. (2008) RNA interference in mammalian cell systems. Curr Top Microbiol Immunol 320, 1–19. 7. Michel, T., Feron, O. (1997) Nitric oxide synthases: which, where, how, and why? J Clin Invest 100, 2146–2152. 8. Kantor, D. B., Lanzrein, M., Stary, S. J., Sandoval, G. M., Smith, W. B., Sullivan,

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B. M., et al. (1996) A role for endothelial NO synthase in LTP revealed by adenovirusmediated inhibition and rescue. Science 274, 1744–1748. Liu, J., Sessa, W. C. (1994) Identification of covalently bound amino-terminal myristic acid in endothelial nitric oxide synthase. J Biol Chem 269, 11691–11694. Busconi, L., Michel, T. (1993) Endothelial nitric oxide synthase. N-terminal myristoylation determines subcellular localization. J Biol Chem 268, 8410–8413. Harding, T. C., Geddes, B. J., Noel, J. D., Murphy, D., Uney, J. B. (1997) Tetracyclineregulated transgene expression in hippocampal neurones following transfection with adenoviral vectors. J Neurochem 69, 2620–2623. Harding, T. C., Geddes, B. J., Murphy, D., Knight, D., Uney, J. B. (1998) Switching transgene expression in the brain using an adenoviral tetracycline-regulatable system. Nat Biotechnol 16, 553–555. Wang, S., Teschemacher, A. G., Paton, J. F., Kasparov, S. (2006) Mechanism of nitric oxide action on inhibitory GABAergic signaling within the nucleus tractus solitarii. FASEB J 20, 1537–1539.

NOS Antagonism Using Viral Vectors as an Experimental Strategy 14. Waki, H., Murphy, D., Yao, S. T., Kasparov, S., Paton, J. F. (2006) Endothelial NO synthase activity in nucleus tractus solitarii contributes to hypertension in spontaneously hypertensive rats. Hypertension 48, 644–650. 15. Waki, H., Kasparov, S., Wong, L. F., Murphy, D., Shimizu, T., Paton, J. F. (2003) Chronic inhibition of endothelial nitric oxide synthase activity in nucleus tractus solitarii enhances baroreceptor reflex in conscious rats. J Physiol 546, 233–242. 16. Paton, J. F., Waki, H., Abdala, A. P., Dickinson, J.„ Kasparov, S. (2007) Vascular-brain signaling in hypertension: role of angiotensin II and nitric oxide. Curr Hypertens Rep 9, 242–247. 17. Bridge, P. M., Ball, D. J., Mackinnon, S. E., Nakao, Y., Brandt, K., Hunter, D. A., et al. (1994) Nerve crush injuries – a model for axonotmesis. Exp Neurol 127, 284–290. 18. Verdu, E., Ceballos, D., Vilches, J. J., Navarro, X. (2000) Influence of aging on peripheral nerve function and regeneration. J Peripher Nerv Syst 5, 191–208. 19. Gonzalez-Hernandez, T., Rustioni, A. (1999) Expression of three forms of nitric oxide synthase in peripheral nerve regeneration. J Neurosci Res 55, 198–207. 20. Sunico, C. R., Portillo, F., Gonzalez-Forero, D., Kasparov, S.„ Moreno-Lopez, B. (2008) Evidence for a detrimental role of nitric oxide synthesized by endothelial nitric oxide synthase after peripheral nerve injury. Neuroscience 157, 40–51. 21. Stegmeier, F., Hu, G., Rickles, R. J., Hannon, G. J., Elledge, S. J. (2005) A lentiviral microRNA-based system for single-copy polymerase II-regulated RNA interference in mammalian cells. Proc Natl Acad Sci USA 102, 13212–13217. 22. Liu, B., Paton, J. F., Kasparov, S. (2008) Viral vectors based on bidirectional cell-

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specific mammalian promoters and transcriptional amplification strategy for use in vitro and in vivo. BMC Biotechnol 8, 49–57. Sunico, C. R., Portillo, F., GonzalezForero, D., Moreno-Lopez, B. (2005) Nitric-oxide-directed synaptic remodeling in the adult mammal CNS. J Neurosci 25, 1448–1458. Coleman, J. E., Huentelman, M. J., Kasparov, S., Metcalfe, B. L., Paton, J. F., Katovich, M. J., et al. (2003) Efficient largescale production and concentration of HIV1-based lentiviral vectors for use in vivo. Physiol Genomics 12, 221–228. Bewig, B., Schmidt, W. E. (2000) Accelerated titering of adenoviruses. Biotechniques 28, 870–873. Lonergan, T., Teschemacher, A. G., Hwang, D. Y., Kim, K. S., Pickering, A. E., Kasparov, S. (2005) Targeting brain stem centers of cardiovascular control using adenoviral vectors: impact of promoters on transgene expression. Physiol Genomics 20, 165–172. Gonzalez-Forero, D., Portillo, F., Sunico, C. R., Moreno-Lopez, B. (2004) Nerve injury reduces responses of hypoglossal motoneurones to baseline and chemoreceptormodulated inspiratory drive in the adult rat. J Physiol 557, 991–1011. Xu, H., McCann, M., Zhang, Z., Posner, G. H., Bingham, V., El-Tanani, M., et al. (2009) Vitamin D receptor modulates the neoplastic phenotype through antagonistic growth regulatory signals. Mol Carcinog 48, 758–772. Chen, X., Xu, H., Wan, C., McCaigue, M., Li, G. (2006) Bioreactor expansion of human adult bone marrow-derived mesenchymal stem cells. Stem Cells 24, 2052–2059. Paxinos, G., Watson, C. (1986) The Rat Brain in Stereotaxic Coordinates. Academic, London.

INDEX

A

Enhanced green fluorescent protein (eGFP) . . . . . . 198, 201 Entrapment efficiency . . . . . . . . . . . . . . . . 172, 175–176, 183

Acidic iodide bath . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82–86 Adenoviral vector (AVV) . . . . . . . . . 154, 198–202, 204, 209 Alkaline single cell gel electrophoresis . . . . . . . 18–19, 22–23 Apollo 4000 free radical detector . . . . . . . . . . . . . . . . . . . . 189 ASHI mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60, 67–69 Astigmatic mirror cells . . . . . . . . . . . . . . . . . . . . . . . . . 118, 127 Atomic force microscopy (AFM) . . . . . . . . . . . . . . . 175, 187

Faraday modulation spectroscopy. . . . . . . . . . . . . . . .123–124 Flow injection analysis (FIA) . . . . . . . . . . . . 91–93, 101–102 Fluorescence . . . . . . . . . . . . . 21, 23, 28, 60, 62–69, 200–201 293FT cells . . . . . . . . . . . . . . . . . . . . . . . . . . 156, 159, 162–163

B

G

Balb/c . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59–60, 67, 70 Bioimaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57–70 BLOCK-iT lentiviral RNAi expression system . . . 155, 158 Blood collection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31–33 Bregma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75–76

GalN . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60, 65, 67, 69 Gene therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8, 154 Griess . . . . . . . . . . . . . . . . . . . . . . . 28, 157, 159, 165–166, 190 Griess Assay . . . . . . . . . . . . . . . . . . . . 157, 159, 165–166, 190 GRSNO . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30 GSNO . . . . . . . . . 29–32, 170–171, 174, 178–181, 183–184 GSNO films . . . . . . . . . . . . . . . . . . . . . . . . . 174, 178–181, 184

C Carboxymethylcellulose (CMC) . . . . . . . . . . . . 189, 192–193 Cardiac ischemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 Cavity enhanced spectroscopy . . . . . . . . . . . . . . . . . . 119–121 C57BL/6 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70 β Cell . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153–167 Cellular oxidative stress . . . . . . . . . . . . . . . . . . . . 158, 166–167 Cell viability . . . . . . . . . . . . . . . . . . . . . . . . . 157, 164–165, 181 Cerebral blood flow (CBF) . . . . . . . . . . . . . . . . . . . . . 7, 75–77 Chemiluminescence. . . . . . . . .27–37, 40–51, 55, 81, 91–92, 94–96, 99, 115, 121, 155, 166, 218–219 Chitosan . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189–191, 194 CHNI . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58–65, 67–70 CL-flow cell . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 92 Compound muscle action potential (CMAP) . . . . . 200–202 Coomassie brilliant blue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67 Cu(II)-ascorbic acid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27 Cu(II)Cl2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28 Cytomegalovirus (CMV) . . . . . . . . . . . . . . . . . . . . . . 198, 200

D Deproteination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 44–45, 52 Diabetes mellitus (T1DM) . . . . . . . . . . . . . . . . . . . . . . . . . . 153 Diethyldithiocarbamate (DETC) . . . . . . . . . . . 135–137, 144 Differential scanning calorimetry (DSC) . . . . 177, 179, 184 Dimethyl sulfoxide (DMSO) . . . . 59, 61, 63, 65, 67, 94, 96, 100, 157, 164 Distributed-feedback (DFB) . . . . . . . . . . . . . . . . . . . . . . . . 117 D-NAME . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84–85, 87–88 Double emulsion methods . . . . . . . . . . . . 172, 175–176, 183

E Electron paramagnetic resonance . . . . . . . . . . . . . . . . . . . . . 74 Endothelium-derived relaxing factor . . . . . . . . . . 3, 6, 57, 73

F

H Hamilton syringe . . . . . . . . 33, 35, 43, 46, 55, 206, 211–212 γ-H2AX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17–24 HEK 293 cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 204, 209 Helmholtz resonant cell . . . . . . . . . . . . . . . . . . . . . . . . . . . . 130 Hepatic Injury Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 Herriott cell. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .118, 127 Hippocampus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73–80 HPLC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45–46, 64–65, 79 HUVECs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5, 70, 83, 85, 87

I I- κB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 154 ImageJ software . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66 Immunofluorescence microscopy . . . . . . . . . . . . . . . . . . 18–19 Inlet. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105–112 INOS . . . . . . . 2, 5–6, 59, 66–67, 74–75, 153–167, 198, 201 Ischemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73–80, 141–142 Isoket . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137, 140–141, 146 ISO-NOPF200 NO probe . . . . . . . . . . . . . . . . . . . . . . . . . . . 74 ISO-NOP sensor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189–190 I3 tri-iodide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42

K Kupffer cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67, 69–70

L Laser absorption spectroscopy (LAS) . . . 116, 118, 123, 126 Laser Doppler Flowmetry (LDF) . . . . . . . . . . . . . 75–76, 182 Lentiviral vector. . . . . . . . . . . . . 153–167, 206–207, 212–215 Lenti-X 293T . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 212–214

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

225

Nitric Oxide

226 Index

Ligation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155, 160 Light mineral oil . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 170, 172 Lipopolysaccharide (LPS) . . . . . . . . . . . . . . 59–60, 64–67, 69 Liquid software . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48, 53–54 L-NAME . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84–85, 87–88 Loading capacity (LC) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 180 Loading efficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 178–180 Luminol . . . . . . . . . . . . . . . . . . . . . . . . . 92–100, 102, 208, 218

M MAHMA NONOate . . . . . . . . . . . . . . . . . . . . . . . . . 108–110 MALDI-TOF. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62–63 Manganese . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59–60 Membrane inlet . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105–112 Membrane inlet mass spectrometry . . . . . . . . . . . . . . 105–112 MGD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 136–146 Microdialysis . . . . . . . 93, 137–138, 140–141, 143, 145–146 Milli-Q . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41, 49, 55, 59 MNIC-MGD . . . . . . . . . . . . . . . . . . . . . . . . . . . . 136, 139–146 MNIP . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58–62, 64–67, 69–70 MTT cell proliferation kit . . . . . . . . . . . . . . . . . . . . . . . . 1, 164

N NF-κB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 154 7-NI . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75, 77, 79 Nitrate . . . . . . . . . . . . . . . . . . . . . 42, 48, 50–51, 74, 82–84, 86 Nitric oxide synthase (eNOS) . . . 2–5, 8, 17, 66, 74, 83, 88, 154, 198–204 Nitrite . . . . . . . . 3, 28–30, 32–33, 35–37, 39–55, 82–86, 92, 95, 102, 108, 110–111, 159, 165–166, 189–191, 193 NONOate. . . . . . . . .108–110, 170–172, 175–178, 181–184 NOS1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159 NOS2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159 NOS3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83, 85, 87–88, 159 Nucleus tractus solitarii (NTS) . . . . . . . . . . . . . . . . . 198–199, 201–204, 209

O Optical parametric oscillators (OPO) . . . . . . . . . . . . . . . . 116 Origin . . . . . . . . . . . . . . . . . . 8, 29, 31, 33–34, 36, 47–48, 201 Oxidation . . . . . . . . . . . . . . . . . . . . . . . . 27–28, 36, 39–55, 138

P Pancreatic islet cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153 PENTR/U6 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155, 158–161 Peroxynitrite . . . . . . . . . . . . . . . . . . . . . . . . 5, 92, 94–95, 97–98 Photoacoustic spectroscopy . . . . . . . . . . . . . . . . 121–123, 125 Plasma . . . . . . . . 28, 32–37, 40–41, 44–45, 51–52, 119, 144 PLGA microparticles . . . . . . . . . . . . . . . . . . . . . 171, 175–178 PTIO . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66

Q Quantum cascade lasers . . . . . . . . . . . . . . . . . . . . . . . . 115–131

R Raw264.7 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7, 60, 64–65, 264 Reaction vessel . . . . . 28–29, 42–43, 45–48, 51–55, 107–110 Reactive nitrogen species (RNS) . . . . . . . . . . . . . . 60, 91–103 Reactive oxygen species (ROS) . . . . 4, 60, 92, 141–142, 154 Recombinant IL-1β . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 157 Red blood cells . . . . . . . . . . . . . . . . . . . . . . . . . . . 28, 32, 34, 40 Reperfusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73–79, 141–142 RIN-r cells . . . . . . . . . . . . . . . . . . . . . . . . . . 157, 159, 164–166 RNA . . . . . . . . . . . . . . . . . . 59, 66–67, 83, 154, 198, 203, 205 RSNO . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27–37

S Scanning electron microscope (SEM) . . . 87, 140, 172, 174, 184 Scavenging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91–103 Sephadex column . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 Sequential Injection . . . . . . . . . . . . . . . . . . 91, 96, 98–99, 101 Sequential injection analysis (SIA) . . . . . . . . . . . . . 91, 93–94 Sievers Nitric Oxide Analyzer . . . . . . . . . . . . . . . . . . . . . . . . 29 Silastic inlet . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 112 Single emulsion methods . . . . . . . . . 171–172, 175–176, 183 SiRNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83–85, 87–88 S-nitrosoglutathione . . . . . . . . . . . . . . . . . . . 29, 170, 174, 183 S-nitroso-N-acetyl–DL-penicillamine (SNAP) . . . . . . . . 75, 78–79 S-Nitrosothiols . . . . . . . . . . . . . . . . . . . . 27–37, 143, 145–146 Sol-Gel. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .190, 193 Spin trapping . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 74, 135–146 Spontaneously hypertensive rat (SHR) . . . . . . . . . . 199, 201, 203–204 Superfine structure (SFS) . . . . . . . . . . . . . . . . . . . . . . . . . . . 141 SYN . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 200, 203

T Teflon . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53, 55, 107, 111 Teflon septum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53, 55 TeNOS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 198–202 Tetramethyl Orthosilicate (TMOS) . . . . . . . . 188–189, 191, 193–194 Thiols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27–28, 36–37, 144 TNFα . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 154 Transduction. . . .6, 157, 159, 163–165, 167, 198, 205–206, 211–212, 215 Transformation . . . . . . . . . . . . . . . . . . . . . . . . . 8, 155, 160–161 TriPure isolation reagent kit . . . . . . . . . . . . . . . . . . . . . . 59, 66

V Viral vectors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205–206

W 1400 W . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75, 77, 79 Western Blotting . . . . . . . . . . . . . . . 84, 86–88, 200, 216–219 White cell . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 127