Hydrogen peroxide as a potential biomarker of oxidative stress is there a reliable assay

HYDROGEN PEROXIDE AS A POTENTIAL BIOMARKER OF OXIDATIVE STRESS: IS THERE A RELIABLE ASSAY? MOHAMED SAH REDHA BIN HAMZAH B.Sc.(Hons.) in Chemistry A THESIS SUBMITTED FOR THE DEGREE OF MASTER OF SCIENCE DEPARTMENT OF BIOCHEMISTRY NATIONAL UNIVERSITY OF SINGAPORE 2007 ACKNOWLEDGEMENTS I would like to convey my deepest and most sincere appreciation to the following people from the NUS Department of Biochemistry: Professor Barry Halliwell for his great patience and useful guidance throughout my project despite his hectic schedule; and most importantly, for providing me with the golden opportunity to be part of his research team; Ms. Long Lee Hua for providing me with the necessary resources; Dr Tang Soon Yew for his valuable opinions and generous sharing of knowledge; Assist. Prof. Andrew Jenner for being approachable for advice; Dr Jan Gruber, Sherry Huang, Wang Huansong, Mary Ng Pei Ern, Chu Siew Hwa, Siau Jia Ling and Li Lingzhi, for their precious contributions to the project; and Prof. Sit Kim Ping and Dr Jetty Lee for their cheerful smiles. I would like to present this work to my parents, and thank them for their love and encouragement. To Andrew Tan Kong Hui, thanks for your support too! Through this journey with the Oxidants and Antioxidants Group, I have cultivated the habit of including fruits and vegetables in my previously unbalanced diet. And I have found out that 100% atmospheric oxygen is not going to help me be a better athlete. i TABLE OF CONTENTS Page Acknowledgements i Table of contents ii Abstract vi List of tables viii List of figures x List of abbreviations and keywords xii CHAPTER 1. INTRODUCTION 1.1. Free Radicals And Reactive Species 1 1.2. Reactive Oxygen Species: Formation 2 1.3. The Good Side Of Reactive Oxygen Species 6 1.4. Antioxidant Defences 8 1.5. Oxidative Stress: The Bad Side Of Reactive Oxygen Species 9 1.6. Use Of Biomarkers In Oxidative Stress Measurement 10 1.7. Hydrogen Peroxide As A Biomarker Of Oxidative Stress 14 1.8. Potential Problems In Hydrogen Peroxide Measurement 16 1.9. Importance Of A Good Analytical Technique 17 1.10. Objectives Of Present Study 18 CHAPTER 2. EXPERIMENTAL PROCEDURES 2.1. Materials ii 2.1.1. Reagents and instrumentation 19 2.1.2. Human subjects 20 2.1.3. Preparation of beverages 21 2.2. Methods 2.2.1. Preparation of hydrogen peroxide standards 21 2.2.2. Preparation of human subjects 22 2.2.3. Oxygen electrode assay 22 2.2.4. Recovery study for oxygen electrode assay 23 2.2.5. Study of ascorbate effect on oxygen electrode assay 24 2.2.6. Ferrous ion oxidation- xylenol orange version 2 (FOX-2) assay 25 2.2.7. Recovery study for FOX-2 assay 26 2.2.8. FeTMPyP-catalysed indamine dye formation assay (FeTMPyP assay) 26 2.2.9. Recovery study for FeTMPyP assay 28 2.2.10. Pentafluorobenzenesulfonyl fluorescein (PFBSF) assay 28 2.2.11. Homovanillic acid (HVA) assay 29 2.2.12. p-Hydroxyphenyl acetic acid (HPAA) assay 29 2.2.13. 2,2’-Azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) assay [ABTS assay] 30 2.2.14. Preformation of ABTS+• and the quenching effect of urine 30 2.2.15. N-Acetyl-3,7-dihydroxyphenoxazine (amplex red) assay 31 2.2.16. Recovery study for amplex red assay 32 2.2.17. 2’,7’-Dichlorodihydrofluorescein (DCFH) assay 33 2.2.18. Recovery study for 2’,7’-dichlorodihydrofluorescein (DCFH) assay 34 iii 2.2.19. Monitoring the progress of DCFH assay and the effect of catalase and SOD 34 2.2.20. Dihydrorhodamine 123 (DHR) assay 35 2.2.21. Recovery study for dihydrorhodamine 123 (DHR) assay 36 2.2.22. Basal urinary hydrogen peroxide measurements in human subjects 36 2.2.23. Coffee drinking study 36 2.2.24. Creatinine assay 37 CHAPTER 3. RESULTS 3.1. Catalase-Based Electrochemical Method 3.1.1. Oxygen electrode assay 38 3.2. Non-Enzymatic Chemical-Based Methods 3.2.1. Ferrous ion oxidation- xylenol orange version 2 (FOX-2) assay 46 3.2.2. FeTMPyP-catalysed indamine dye formation assay (FeTMPyP assay) 57 3.2.3. Pentafluorobenzenesulfonyl fluorescein (PFBSF) assay 65 3.3. Peroxidase-Based Methods 68 3.3.1. Homovanillic acid (HVA) assay 69 3.3.2. p-Hydroxyphenyl acetic acid (HPAA) assay 73 3.3.3. 2,2’-Azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) assay [ABTS assay] 76 3.3.4. N-Acetyl-3,7-dihydroxyphenoxazine (amplex red) assay 83 3.3.5. 2’,7’-Dichlorodihydrofluorescein (DCFH) assay 92 3.3.6. Dihydrorhodamine 123 (DHR) assay 103 3.4. Basal Urinary Hydrogen Peroxide Measurements 109 iv 3.5. Effect Of Coffee On Basal Urinary Hydrogen Peroxide 112 CHAPTER 4. FURTHER DISCUSSION 114 CHAPTER 5. CONCLUSION 131 CHAPTER 6. REFERENCES 132 v ABSTRACT Oxidative stress causes damage to the critical biomolecules in humans. When left unchecked, it contributes to the development of several diseases such as cancer, diabetes, cardiovascular diseases, neurodegenerative disorders and even to the process of ageing itself. Considerable debate over identifying the best biomarkers of oxidative stress is still ongoing. Good biomarkers like F2-isoprostanes have been proposed to be among the most reliable but they require the use of expensive instrumentation and extensive preparation steps. But hydrogen peroxide (H2O2) can be easily detected in freshly-voided human urine, without the need for costly set-ups, and it has been proposed as a biomarker of oxidative stress. Obtaining urine also does not require an invasive sampling procedure. In order to investigate how well H2O2 fits into the criteria of an ideal biomarker, an assay that is highly specific, sensitive and reproducible for urinary H2O2 measurement is first required. In the present study, current methods of H2O2 measurement in urine samples (by FOX-2 and oxygen electrode assays) were examined, and various other peroxidasebased and non-enzymatic, chemical-based methods were developed and tested for their suitability to measure H2O2 in urine. The classical oxygen electrode assay and the newlydeveloped, DCFH peroxidase-based assay emerged to be the two most reliable methods. The DCFH assay was able to detect a basal level of H2O2 excreted by healthy individuals, with less intra-individual variation throughout the day and between different days than with the oxygen electrode assay. In future, urinary H2O2 can be further studied with the DCFH assay, alongside other classical biomarkers of oxidative stress, in known pathological conditions and to see the effect of intervention of these conditions with vi antioxidant therapy. Hence, the importance of a good analytical technique can never be overemphasized; in the study of biomarkers of oxidative stress, any data would be meaningless if the method that generates them is not suitable for that application. vii LIST OF TABLES Page 1.1. Nomenclature of reactive species found in vivo 1 1.2. Biomarkers of oxidative stress/damage associated with some human diseases 14 1.3. Data of urinary hydrogen peroxide analyzed by 3 different ways 16 3.1. Accuracy of determination of PBS solutions of H2O2 by the O2 electrode assay 41 3.2. O2 electrode assay recovery study 42 3.3: First study of ascorbate effect on O2 electrode assay 43 3.4: Second study of ascorbate effect on O2 electrode assay 45 3.5. FOX-2 assay recovery study 48 3.6. Comparison of FOX-2 assay with O2 electrode assay in one individual 50 3.7. Comparison of FOX-2 Assay with O2 electrode assay in a few individuals 51 3.8. Effect of dilution of urine sample on FOX-2 assay 53 3.9. Comparison of 10xD-FOX-2 assay with O2 electrode assay 54 3.10. 10xD-FOX-2 assay recovery study 55 3.11. FeTMPyP assay recovery study 61 3.12. Effect of dilution of urine sample on HVA assay 71 3.13. HVA assay: fluorescence in different mixtures 72 3.14. Effect of dilution of urine sample on HPAA assay 74 3.15. ABTS assay: sample absorbance data 79 3.16. Quenching effect of urine on preformed ABTS+• 80 3.17. Effect of dilution of urine sample on amplex red assay 85 viii 3.18. Amplex red assay recovery study 87 3.19. Comparison of the amplex red assay with O2 electrode assay in a few individuals 88 3.20. Effect of dilution of urine sample on DCFH assay 94 3.21. DCFH assay recovery study 96 3.22. Comparison of the DCFH assay with O2 electrode assay in a few individuals 99 3.23. Coefficient of variation of various urine samples analyzed by DCFH assay 100 3.24. Effect of dilution of urine sample on DHR assay and comparison with DCFH assay 106 3.25. DHR assay recovery study and comparison with DCFH assay 107 3.26. Variations in H2O2 level throughout the day as measured by two assays (DCFH and O2 electrode) 111 3.27. Effect of coffee consumption on urinary H2O2 concentration 113 ix LIST OF FIGURES Page 1.1. Molecular orbital diagram of dioxygen 2 1.2. Pathways of ROS formation, the lipid peroxidation process and the role of glutathione and other antioxidants – Vitamin E, Vitamin C, lipoic acid – in the management of oxidative stress 4 1.3. ROS-induced MAPK signaling pathways 7 1.4. Chemical structure of (a) 8-hydroxy-2’-deoxyguanosine (8OHdG) and (b) 8iso-Prostaglandin F2α 12 3.1. O2 electrode chart recording 38 3.2. A standard calibration plot for the O2 electrode assay 40 3.3. A standard calibration plot for the FOX-2 assay 47 3.4. Structures of (a) hemin and (b) FeTMPyPCl5 57 3.5. Coupling reaction to form indamine dye 58 3.6. Absorbance progress of the FeTMPyP-catalyzed indamine dye formation reaction 58 3.7. A standard calibration plot for the FeTMPyP assay 60 3.8. FeTMPyP reaction scheme 63 3.9. Pentafluorobenzenesulfonyl fluorescein (PFBSF) 65 3.10. Oxidation of HVA in the presence of HRP to a fluorescence dimer 69 3.11. A standard calibration plot for the HVA assay 70 3.12. A standard calibration plot for the HPAA assay 74 3.13. Structure of ABTS and its oxidation products 76 3.14. A standard calibration plot for the ABTS assay 78 3.15. Chemical structures of polyphenols and their metabolites detected in urine 81 x 3.16. Mechanism of action for (a) ABTS/HRP/ H2O2 and (b) ascorbic acid (AA) with ABTS+• 82 3.17. A standard calibration plot for amplex red assay 84 3.18. HRP-catalyzed amplex red oxidation by H2O2 89 3.19. Structures of some peroxidase substrates 91 3.20. A standard calibration plot for DCFH assay 93 3.21. Fluorescence intensity progress of the DCFH/HRP reaction with 0 to 10 µM H2O2 standards 97 3.22. Fluorescence intensity progress of the DCFH/HRP reaction in 2 urine samples (S1 & S2) 102 3.23. Oxidation of dihydrorhodamine 123 (DHR) to rhodamine 123 103 3.24. A standard calibration plot for DHR assay 104 4.1. Mechanism of DCFH-DA de-esterification to DCFH and further oxidation to highly-fluorescent DCF by ROS and RNS 121 4.2. Schematic representation of DCFH oxidation by HRP initiated by H2O2 123 4.3. Chemical structure of scopoletin 126 4.4. Comparison of structures of HFLUOR (dihydrofluorescein) and DCFH (2’,7’-dichlorodihydrofluorescein), as well as their oxidized products 130 4.5. Molecular structures of ascorbic acid quenchers (AAQs) 130 xi LIST OF ABBREVIATIONS AND KEYWORDS AA Ascorbic acid AAQ Ascorbic acid quencher ABTS 2,2’-Azino-bis(3-ethylbenzothiazoline-6-sulfonic acid AscH- Ascorbate CuZnSOD Copper and zinc-containing superoxide dismutase DCF 2’,7’-Dichlorofluorescein DCFH 2’,7’-Dichlorodihydrofluorescein DCFH-DA 2’,7’-Dichlorodihydrofluorescein diacetate DHA Dehydroascorbic acid DHR Dihydrorhodamine 123 DMSO Dimethyl sulfoxide EC-SOD Extracellular superoxide dismutase FeTMPyP meso-Tetrakis(1-methyl-4-pyridyl)porphinatoiron(III) FOX Ferrous ion oxidation – xylenol orange (assay) FOX-2 Ferrous ion oxidation – xylenol orange version 2 (assay) 10xD-FOX-2 FOX-2 (assay) conducted on sample(s) diluted by a factor of 10 GC-MS Gas chromatography – mass spectrometry GC-MS/MS Gas chromatography – tandem mass spectrometry GPx Glutathione peroxidase GRed Glutathione reductase GSH Glutathione (reduced form) xii GSSG Glutathione (oxidized form) HEPES 2-[4-(hydroxyethyl)-1-piperazinyl]ethanesulfonic acid HFLUOR Dihydrofluorescein HPAA p-Hydroxyphenyl acetic acid HPLC High performance liquid chromatography HRP Horseradish peroxidase HVA Homovanillic acid or 4-hydroxy-3-methoxy-phenylacetic acid LC-MS Liquid chromatography – mass spectrometry LC-MS/MS Liquid chromatography – tandem mass spectrometry LOOH Lipid hydroperoxide MAPK Mitogen-activated protein kinase (pathway) MDHA Monodehydroascorbic acid MeOH Methanol MnSOD Manganese-containing superoxide dismutase MS Mass spectrometry NAD(P)+ Nicotinamide adenine dinucleotide (phosphate) NAD(P)H Reduced nicotinamide adenine dinucleotide (phosphate) NF-κB Nuclear factor κB PBS Phosphate buffered saline PFBSF Pentafluorobenzenesulfonyl fluorescein RFU Relative fluorescence unit(s) RNS Reactive nitrogen species ROS Reactive oxygen species xiii SOD Superoxide dismutase SDS Sodium dodecylsulfate TEMPO 2,2,6,6-tetramethyl-1-piperidinyloxy (radicals) TNF Tumour necrosis factor T-OH α-Tocopherol or Vitamin E UA Uric acid xiv CHAPTER 1 INTRODUCTION 1.1. FREE RADICALS AND REACTIVE SPECIES What are free radicals? A free radical is defined as any chemical species capable of independent existence (hence, termed ‘free’) that contains one or more unpaired electrons in atomic or molecular orbitals (Halliwell & Gutteridge, 1999). Free radicals and other reactive species are continuously generated in vivo during physiological and pathological processes. Table 1.1 lists some of the reactive species that can be found in vivo. Table 1.1. Nomenclature of reactive species found in vivo (adapted from Halliwell et al., 2004b) REACTIVE SPECIES Nonradicals Free radicals Reactive oxygen species (ROS) Superoxide, O2•Hydrogen peroxide, H2O2 Hydroxyl, OH• Hypobromous acid, HOBr Hydroperoxyl, HO2• Hypochlorous acid, HOCl Ozone, O3 Peroxyl, RO2• Singlet oxygen 1∆g O2 • Alkoxyl, RO Organic peroxides, ROOH Carbonate, CO3•Peroxynitrite, ONOO•Carbon dioxide, CO2 Peroxynitrous acid, ONOOH Reactive nitrogen species (RNS) Nitric oxide, NO• Nitrogen dioxide, NO2• Nitrous acid, HNO2 Nitrosyl cation, NO+ Nitroxyl anion, NODinitrogen tetroxide, N2O4 Dinitrogen trioxide, N2O3 Peroxynitrite, ONOOPeroxynitrous acid, ONOOH Nitronium (nitryl) cation, NO2+ Alkyl peroxynitrites, ROONO Nitryl (nitronium) chloride, NO2Cl 1 1.2. REACTIVE OXYGEN SPECIES: FORMATION . By the given definition of ‘free radical’, molecular oxygen (or dioxygen) in the ground state has an electronic configuration that qualifies it to be a biradical; it has two unpaired electrons with parallel spins, each located in a different π* antibonding orbital (Fig. 1.1). Fig. 1.1. Molecular orbital diagram of dioxygen (obtained from www.steve.gb.com) The presence of unpaired electron(s) in a free radical usually confers it a considerable degree of reactivity and this probably accounts for the reactivity of dioxygen with other radical molecules (Valko et al., 2004). Radicals derived from oxygen represent the most important class of radical species generated in living systems. These oxygencontaining radicals, together with some other non-radical, oxygen-containing 2 molecules/ions, are generally termed as reactive oxygen species (ROS), which together with the reactive nitrogen species (RNS), are products of normal cellular metabolism (Table 1.1). These species are well-recognized for playing a dual role as both deleterious and beneficial species, since they can be either harmful or beneficial to living systems (Valko et al., 2007). The addition of one electron to dioxygen forms the superoxide anion radical (O2•-) (Miller et al., 1990). Its production occurs mostly within the mitochondria due to the ‘leakage’ of a small number of electrons from the electron transport chain which is the main source of ATP in most mammalian cells. O2•- is produced from Complexes I and III located at the inner mitochondrial membrane and released into the matrix as well as the intermembranous space (Camello-Almaraz et al., 2006). O2•- is also produced from the direct reaction of autooxidizable molecules with dioxygen, as well as through the action of certain enzymes such as xanthine oxidase and galactose oxidase (Halliwell & Gutteridge, 1999). O2•- cannot directly attack DNA, proteins or lipids, but at elevated levels, can mobilize small amounts of iron from the iron-storage protein ferritin (Bolann et al., 1990). It can also attack the active sites of some enzymes containing iron-sulphur clusters, causing their inactivation accompanied by iron release (Liochev, 1996). Hydrogen peroxide (H2O2) is produced through the spontaneous or enzymatic dismutation of O2•- (2 O2•- + 2 H+ → H2O2 + O2). H2O2 can also be produced directly by several enzymes such as xanthine oxidase. It is poorly reactive with most biomolecules and appears unable to directly oxidize DNA, lipids and proteins, except for a few proteins which have hyper-reactive thiol groups or methionine residues (Halliwell & Gutteridge, 1999). The danger of H2O2 largely comes from its ready conversion to the 3 Fig. 1.2. Pathways of ROS formation, the lipid (LH) peroxidation process and the role of glutathione (GSH) and other antioxidants – Vitamin E (T-OH), Vitamin C(AscH-), lipoic acid – in the management of oxidative stress (adapted from Valko et al., 2007) 4 indiscriminately reactive hydroxyl radical (OH•), either by exposure to ultraviolet light (H2O2 → 2OH•) or through the Fenton reaction (Halliwell et al., 2000a). Iron released by O2•- (or other transition metal ions) can participate in the Fenton reaction with H2O2 to generate OH• and the reaction can be perpetuated by any reducing agent (e.g. ascorbic acid and O2•-) capable of recycling Fe3+ back to Fe2+ (Halliwell & Gutteridge, 1999): H2O2 + Fe2+ → Fe3+ + OH• + OHFe3+ + O2•- → Fe2+ + O2 With a high level of reactivity and very short half-life of approximately 10-9 s in vivo, OH• reacts close to its site of formation (Valko et al., 2007). OH• can attack and damage all biomolecules: carbohydrates, lipids, proteins and DNA (Von Sonntag, 1987). When lipids are attacked by OH•, the chain reaction of lipid peroxidation starts and lipid hydroperoxides (LOOH) accumulate. These can be degraded in the presence of iron or copper ions (Halliwell & Gutteridge, 1999): LOOH + Fe2+ → Fe3+ + LO• + OHLOOH + Fe3+ → Fe2+ + LOO• + H+ The resulting alkoxyl (LO•) and peroxyl (LOO•) radicals can damage membrane proteins and also attack new lipid molecules to propagate lipid peroxidation. Fig. 1.2 summarizes the various pathways of ROS formation. 5 1.3. THE GOOD SIDE OF REACTIVE OXYGEN SPECIES ROS are known to play a role in several aspects of intracellular signaling and regulation (Valko et al., 2007). Most cell types have been shown to generate low concentrations of ROS which act as secondary messengers in signal transduction cascades when the cell receptors are stimulated by cytokines, growth factors and hormones (Kamata et al., 1999). The most significant effect of ROS on signaling pathways has been observed in the mitogen-activated protein kinase (MAPK) pathways which involve the activation of nuclear transcription factors (Sun et al., 1996). These factors control the expression of protective genes that repair damaged DNA, power the immune system, arrest the proliferation of damaged cells and induce apoptosis. For example, the p53 protein guards a cell-cycle checkpoint, as inactivation of p53 favours uncontrolled cell division and is associated with more than half of all human cancers (Sun et al., 1996). ROS have been implicated as second messengers involved in the activation of nuclear factor NF-κB via tumour necrosis factor (TNF) and interleukin-1 (Hughes et al., 2005). NF-κB regulates several genes involved in cell transformation, proliferation and angiogenesis, and is involved in inflammatory responses (Valko et al., 2007). Fig. 1.3 gives a diagrammatic summary of the activation of MAPK signaling pathways. ROS production by activated neutrophils and macrophages is a vital component of host organism defense; the phagocytic isoform of NADPH oxidase produces O2•- and other ROS that play essential roles in killing many types of bacteria and other invaders (DeCoursey et al., 2005). The conversion of O2 to O2•- transiently increases the O2 consumption of the cell up to 100 fold, hence the misnomer ‘respiratory burst’ (because it 6 is unrelated to mitochondrial respiration), while the concentration of H2O2 may reach a level of 10-100 µM in the inflammatory environment (DeCoursey et al., 2005; Valko et al., 2007). ROS are also involved in other roles such as cell adhesion, redox regulation of immune responses and as a sensor for changes of oxygen concentration (Frein et al., 2005; Waypa et al., 2005; Valko et al., 2007). Fig. 1.3. ROS-induced MAPK signaling pathways (adapted from Valko et al., 2007) 7 1.4. ANTIOXIDANT DEFENCES Exposure to free radicals from a variety of sources has led organisms to evolve an antioxidant defense system comprising the following (Halliwell & Gutteridge, 1999): (a) Agents (enzymes) that catalytically remove free radicals and other ‘reactive species’. Examples are superoxide dismutase (SOD), catalase, peroxidase and ‘thiol specific antioxidants’. (b) Proteins that minimize the availability of pro-oxidants such as iron ions, copper ions and heme. Some examples are protein transferrins that sequester iron so that none exists ‘free’ in plasma, caeruloplasmins that bind to plasma copper, and ferritins and metallothioneins which store excess iron and copper respectively, within cells. (c) Proteins that protect biomolecules against damage (including oxidative damage) by other mechanisms, e.g. heat shock proteins. (d) Low molecular mass agents that scavenge ROS and RNS. Examples are glutathione, α-tocopherol, ascorbic acid, bilirubin and uric acid. SOD helps to diminish the direct damage caused by O2•- by accelerating its dismutation to H2O2. The most important SOD appears to be manganese-containing SOD (MnSOD), which is located in the mitochondrial matrix; transgenic mice lacking this enzyme die soon after birth with severe mitochondrial damage in many tissues (Li et al., 1995). Copper- and zinc-containing SOD (CuZnSOD) is mostly located in the cytosol of animal cells while extracellular SOD (EC-SOD) is found on the cell surface of many tissues (Halliwell & Gutteridge, 1999). 8 H2O2 can be removed by catalase, an exclusively peroxisomal enzyme in most tissues, as well as by glutathione peroxidase (GPx) (Chance et al., 1979): 2 GSH + H2O2 → 2 H2O + GSSG Oxidized glutathione (GSSG) is reduced back to glutathione (GSH) by glutathione reductase (GRed) (Chance et al., 1979): GSSG + NADPH + H+ → 2 GSH + NADP+ Ascorbate (AscH-) and α-tocopherol (T-OH) are derived from the diet; the former can scavenge many reactive species, including O2•-, LO•, LOO•, OH• and ONOO(Halliwell, 1996). T-OH is a powerful chain-breaking antioxidant that inhibits lipid peroxidation by scavenging LOO• (Halliwell & Gutteridge, 1999). Fig. 1.2 shows some reaction pathways of the earlier discussed antioxidants. 1.5. OXIDATIVE STRESS: THE BAD SIDE OF REACTIVE OXYGEN SPECIES Free radicals and reactive species operate at a low, ‘steady-state’ concentration, measurable in cells, determined by the balance between their rates of production and their rates of removal by the antioxidant defence system which was briefly discussed earlier. Oxidative stress occurs when there is a serious disturbance in this pro-oxidant – antioxidant balance in favour of the former, leading to potential damage (Sies, 1991). Oxidative stress can result from (Halliwell et al., 2004b): (a) Diminished levels of antioxidants, which can arise due to mutations affecting activities of antioxidant defence enzymes such as CuZnSOD or GPx, toxins that 9 deplete antioxidant defences (such as the depletion of GSH by high doses of xenobiotics), or deficiencies in dietary minerals and antioxidants (b) Increased production of reactive species, for example through inappropriate activation of phagocytic cells in chronic inflammatory diseases, or exposure to elevated levels of O2 or other toxins that are either reactive species themselves (e.g. NO2•) or are metabolized to generate reactive species (e.g. paraquat) A major consequence of oxidative stress is damage to nucleic acid bases, lipids and proteins, which can severely compromise cell health and viability or induce a variety of cellular responses through generation of secondary reactive species, ultimately leading to cell death by necrosis or apoptosis (Dalle-Donne et al., 2006). It is widely believed that oxidative damage to biomolecules, if left unchecked, contributes to the development of several diseases such as cancer, cardiovascular diseases, diabetes, neurodegenerative disorders and even to the process of ageing itself (Halliwell & Gutteridge, 1999). 1.6. USE OF BIOMARKERS IN OXIDATIVE STRESS MEASUREMENT The localization and effects of oxidative stress, as well as information regarding the nature of the ROS, may be gleaned from the analysis of discrete biomarkers of oxidative stress/damage isolated from tissues and biological fluids. Biomarkers are defined as characteristics that can be objectively measured and evaluated as indicators of normal biological processes, pathogenic processes, or pharmacologic responses to a therapeutic intervention (Dalle-Donne et al., 2006). 10 Several criteria for an ideal biomarker of oxidative stress/damage can be listed. Very importantly, it must first be measurable by a robust method or assay that is specific, sensitive and reproducible for the biomarker of interest, and detectable even in normal, healthy individuals. Its levels shall not vary widely in the same subjects under the same conditions at different times. Ideally, it shall be predictive of the later development of the disease, though no biomarker has fulfilled this criterion as necessary experiments have not been done (Halliwell et al., 2004a). Biomarker stability is also crucial and since most ROS are generally too reactive and/or have a half-life too short (not more than seconds) to allow direct measurements in cells/tissues or body fluids, their more stable oxidation target products are measured instead, for e.g. lipid peroxidation products (Dalle-Donne et al., 2006). The biomarker must be measurable with relatively small within-assay intrasample variation compared with between-person variations. Whether obtaining the biomarker requires an invasive method or not can be an important factor for consideration, especially when critically-ill patients are involved or when frequent sampling is required. Biological samples that have been used in previous studies include blood, plasma, urine, bronchoalveolar lavage fluid, cerebrospinal fluid, synovial fluid and tissue biopsies (Dalle-Donne et al., 2006). An example of a common biomarker is 8-hydroxy-2’-deoxyguanosine (8OHdG; Fig. 1.4a) which is frequently measured as a biomarker of oxidative damage to DNA (Kasai, 1997). Besides the availability of this assay, other factors supporting 8OHdG measurement include (a) its formation in DNA by several reactive species such as OH• and singlet oxygen, (b) its established mutagenicity in inducing GC→TA transversions, and (c) the multiple mechanisms that have evolved to remove 8OHdG from DNA, or to 11 prevent its incorporation into cellular DNA, which suggests that the cell ‘perceives’ 8OHdG to be a threatening lesion that has to be removed rapidly (Kasai, 1997). However, levels of 8OHdG are not a quantitative marker of damage to DNA by all reactive species (for example, 8OHdG is only a minor product of attack by RNS), and ROS attack on guanine residues yield not only 8OHdG, but also products such as Fapyguanine whose amount relative to that of 8OHdG depends on the redox state of the cell and the presence of transition metal ions (Halliwell, 2000b). Hence, the same amount of free radical attack on DNA can give different levels of 8OHdG. Another drawback is the artifactual generation of 8OHdG during DNA isolation from tissues, hydrolysis and analysis. Consideration should also be given to other DNA base damage products which are known to be mutagenic, quantitatively more important and less ready to form artifactually than 8OHdG (Halliwell, 2000b). Nevertheless, 8OHdG is not readily metabolized and urinary 8OHdG is not confounded by diet (Cooke et al., 2005). Fig. 1.4. Chemical structure of (a) 8-hydroxy-2’-deoxyguanosine (8OHdG) and (b) 8-isoProstaglandin F2α. (b) is the most thoroughly investigated F2-isoprostane. 12 At present, measurement of the biomarker F2-isoprostanes (Fig. 1.4b) is regarded as the most reliable approach to assess free radical-mediated lipid peroxidation in vivo (Montuschi et al., 2004). They are produced from the free radical-induced peroxidation of arachidonic acid esterified to phospholipids (Morrow et al., 1990). Available data indicate that their quantification in either plasma or urine gives a highly precise and accurate index of oxidative stress (Morrow, 2005). They are stable in isolated samples of body fluids, like urine and exhaled breath condensates, providing a non-invasive route for their measurements (Dalle-Donne et al., 2006). Their measured values do not exhibit diurnal variations and are not affected by lipid content in the diet (Richelle et al., 1999). However, F2-isoprostanes have been only reliably measured using mainly mass spectrometric-based (MS-based) methods such as gas chromatography-mass spectrometry (GC-MS) and liquid chromatography-mass spectrometry (LC-MS) methods, and tandem MS methods with either LC or GC (Lee et al., 2004; Liang et al., 2003). Though F2-isoprostanes can be measured accurately down to picomolar concentrations with these methods, the instrumentations involved are expensive; moreover, extensive sample preparation and clean-up (e.g. phospholipid extraction, alkaline hydrolysis and derivatization) are required while great care must be taken to avoid any artifactual formation during this long processing as well as during sample storage (Dalle-Donne et al., 2006). Considerable debate over identifying the best biomarkers of oxidative stress is still ongoing and Table 1.2 shows many other commonly-used biomarkers of oxidative stress/damage and the diseases with which they are associated. 13 Table 1.2. Biomarkers of oxidative stress/damage associated with some human diseases (adapted from Valko et al., 2007). NO2-Tyr, 3-nitrotyrosine 1.7. HYDROGEN PEROXIDE AS A BIOMARKER OF OXIDATIVE STRESS As mentioned earlier, H2O2 plays an important role as an inter- and intra-cellular signaling molecule, so a basal level of H2O2 must be present. In fact, levels of H2O2 at or below about 20-50 µM seem to have limited cytotoxicity to many cell types, while levels above 50 µM have been described as cytotoxic to a wide range of cultured animal, plant and bacterial cells (Halliwell et al., 2000a). H2O2 has been detected in human exhaled breath condensates and the amounts of exhaled H2O2 appear greater in subjects with inflammatory lung diseases (Rosias et al., 2006) and in cigarette smokers (Nowak et al., 2001). H2O2 is also present in the aqueous 14 humor, probably due to the oxidation of ascorbic acid which is normally present in high concentration in these fluids (Reddy, 1990). Oxidative damage to the ocular lens leading to cataract is slowed down by the presence of antioxidant defenses like glutathione (Lou, 2003). On the other hand, H2O2 is low or almost zero in human blood plasma (Frei et al., 1988), likely due to its reaction with heme proteins, ascorbate and protein thiol groups, or metabolism after diffusion into erythrocytes or other cells. The excretion of hydrogen peroxide in human urine was demonstrated for the first time by Varma et al. (1990). Since then, many laboratories have confirmed the presence of significant amounts of H2O2 in freshly-voided urine (Kuge et al., 1999; Long et al., 1999b & 2000; Hiramoto et al., 2002). Thus, it was wondered if H2O2 levels in urine might be a simple biomarker of oxidative stress. While a lot of good has been said of F2isoprostanes, and urinary/plasma o,o’-dityrosine and 3-nitrotyrosine being promising biomarkers to be worked on (Dalle-Donne et al., 2006), costly tandem MS methods (GCMS/MS and LC-MS/MS) are the recommended instrumentations. However, H2O2 can be easily measured in urine without the need for expensive techniques like MS or electron spin-resonance spectroscopy, and in a shorter period of time (Long et al., 1999b). Thus, the possibility that urinary H2O2 is a biomarker of the extent of whole body oxidative stress is a very attractive concept to test (Yuen et al., 2003), and if the results are positive, oxidative stress assessment could be easily done by laboratories of any scale. 15 1.8. POTENTIAL PROBLEMS IN HYDROGEN PEROXIDE MEASUREMENT The rationale for conducting the present study arose when urine samples from one human subject were analyzed for hydrogen peroxide. The subject was a healthy nonsmoker who consumed a dietary supplement pill (GNC’s MegaMen) every morning. Five different samples were collected from him at the stated times (Table 1.3) within a day and were immediately analyzed by three assays, namely the oxygen electrode assay, the ferrous ion oxidation- xylenol orange version 2 (FOX-2) assay and the fluorescence assay. The procedure used for the first two assays could be found in Chapter 2. The fluorescence assay was attempted using the amplex red-peroxidase assay kit (A22188) from Molecular Probes, Inc., and following protocol provided by the supplier company. Table 1.3. Data of urinary hydrogen peroxide analyzed by 3 different ways. One Day / Subject X Concentration of H2O2 in µM of urine samples Sample collection times 1100 hrs 1215 hrs 1330 hrs 1445 hrs 1600 hrs FOX-2 assay 11.7 12.8 13.4 16.4 8.86 A22188 assay 2.56 3.05 3.09 1.91 3.25 O2 Electrode assay 31.1 34.8 46.3 44.5 33.5 The data, as given in Table 1.3, show that for every collection, there was a significant intra-sample variation between the 3 assays. The A22188 assay gave the lowest urinary H2O2 concentration values at all times while the O2 electrode assay gave the largest values. Although the FOX-2 and O2 electrode assay gave values which 16 differed considerably in magnitude, a similar trend of increase and decrease in H2O2 concentrations from 1100hrs to 1600hrs was observed between the two assays. This finding led us to many questions. If these established assays have been so widely used in various types of work, why are they giving very different values of the analyte in the same sample? Which of these methods is giving the correct (or wrong) value? Or, is it possible that none of the methods is giving the right value? If these assays are not valid for urinary H2O2 measurements in the first place, can they be better tailored to meet the specific needs of the study? Are there constituents in urine (such as excreted metabolites from the pill or diet) that can affect accurate measurements of H2O2 by these assays? Is there any chance that these interfering constituents can be identified and/or removed from urine samples which are already biochemically complex to begin with? Or, better yet, are there any other more suitable assays that can be developed to measure urinary H2O2 concentration accurately and hence be able to determine if H2O2 excreted in urine can be a suitable biomarker of oxidative stress? 1.9. IMPORTANCE OF A GOOD ANALYTICAL TECHNIQUE H2O2 is one of the most stable ROS (O2•-, OH•, and singlet oxygen have much shorter life time), offering the opportunity to carefully quantitate the production of a ROS by biological systems (Votyakova et al., 2004). However the accuracy of such determinations also depends on the specificity of the assay system. In the present study, I attempt to answer as many of the questions that were raised at the end of the previous section as possible. The bottom line is that a good analytical 17 technique is required to measure urinary H2O2 accurately before we can confidently say whether it has the potential to be an excellent biomarker of the extent of whole body oxidative stress or not. 1.10. OBJECTIVES OF PRESENT STUDY In summary, the objectives of the present study are to: (a) analytically validate the current methods of hydrogen peroxide measurement in human urine samples (FOX-2 and O2 electrode assays) used in our laboratory and elsewhere; (b) develop a new assay suitable for the measurement of urinary H2O2 that is simple, accurate, sensitive, specific, reproducible and robust; and (c) use the assay developed in (b) to investigate if urinary H2O2 can meet as many of the requirements set out for an ideal biomarker of oxidative stress as possible. 18 CHAPTER 2 EXPERIMENTAL PROCEDURES 2.1. MATERIALS 2.1.1. Reagents and instrumentation All chemicals were of the highest grade available from the stated companies: Hydrogen peroxide (H2O2; 30-35%) from Kanto Chemical Co. Inc., Japan; phosphate buffered saline (PBS; 8 g/L NaCl, 0.2 g/L KCl, 1.44 g/L Na2HPO4 and 0.24 g/L KH2PO4; pH 7.4) from the National University Medical Institute, Singapore (NUMI); 2-[4(hydroxyethyl)-1-piperazinyl]ethanesulfonic acid (HEPES; 99.5% by titration) from Sigma; methanol (MeOH; HPLC grade) from Fisher Scientific; hydrochloric acid (HCl; 37% fuming) from Merck; sulfuric acid (H2SO4; min. 98%) from Merck; N,Ndimethylaniline (min. 99.5%, purified by re-distillation) from Aldrich; dimethyl sulfoxide (DMSO; min. 99.5%, cell culture grade) from AppliChem, Germany; ethanol (min. 99.7%) from BDH AnalaR; glacial acetic acid from JT Baker; sodium hydroxide (NaOH) pellets from Merck; phosphoric acid (H3PO4; 85 %) from Mallinckrodt; 10% SDS solution from Invitrogen; 3-methyl-2-benzothiazolinone hydrazone hydrochloride monohydrate from Fluka; meso-tetrakis(1-methyl-4-pyridyl)porphinatoiron(III) pentachloride (FeTMPyPCl5) from Cayman Chemicals; potassium chloride (KCl) from BDH AnalaR; disodium hydrogen phosphate (Na2HPO4) from Merck; picric acid (1% solution in water) from Aldrich; boric acid from Sigma; creatinine standard (3.0 mg/dl) from Sigma; L-ascorbate, sodium salt (98%, powder) from Sigma; ferrous ammonium 19 sulfate from Sigma; xylenol orange from Sigma; butylated hydroxytoluene from Sigma; pentafluorobenzenesulfonyl fluorescein (PFBSF) from Calbiochem; homovanillic acid (HVA) from Sigma; amplex red (5 mg) from Invitrogen; p-hydroxyphenylacetic acid (HPAA) from Aldrich; 2’,7’-dichlorodihydrofluorescein diacetate (DCFH-DA; 50 mg) from Axxora Platform; 2,2’-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) or ABTS from Sigma; dihydrorhodamine 123 (DHR; 10 mg) from Cayman Chemicals; catalase (EC 1.11.1.6; type C40 from bovine liver; lyophilized, 16,400 U/mg protein) from Sigma; superoxide dismutase (SOD; EC 1.15.1.1; from bovine erythrocyte; copper and zinc-containing i.e. CuZnSOD; lyophilized powder, 3700 U/mg solid) from Sigma; horseradish peroxidase (HRP; EC 1.11.1.7; lyophilized powder, 1067 U/mg solid) from Fluka; and water (MilliQ ultrapure of at least 18.2 MΩ). Instruments used include the Molecular Devices Spectra MAX Gemini EM (for fluorescence readings), Beckman DU 640B spectrophotometer (for UV/Visible absorbance measurements) and a Hansatech oxygen electrode. 2.1.2. Human subjects Healthy men and women aged 19 to 43 years were recruited from the Department of Biochemistry, National University of Singapore. All subjects had Body Mass Index (BMI) within the range of 17 to 24 (i.e. no overweight or obese subjects were recruited) and had no history of cancer, hypertension, diabetes, cardiovascular or liver diseases. Subjects were recruited regardless of race and gender. All subjects were non-smokers, non-vegetarians and were not regular coffee drinkers, except for two subjects who were told to abstain from coffee for at least 14 hours before an experiment. Recruited subjects 20 were not taking any form of oral medications or nutritional supplementations during the period of study (one of the subjects who took a dietary supplement pill daily stopped consuming them for at least 48 hours before an experiment). Subjects were briefed on the procedures and requirements of the study. All subjects gave informed consent. 2.1.3. Preparation of beverages Nestlé ‘Original Ice Coffee’, a typical coffee drink containing milk and sugar, manufactured in Malaysia and packed in 240-ml cans, was bought from a local convenience store and served to subjects chilled. The product was chosen mainly because the consistency of the drinks’ contents was assured by the quality system of Nestlé and self-making of coffee in the kitchen might not be as reliable for repeat experiments. 2.2. METHODS 2.2.1. Preparation of hydrogen peroxide standards A stock concentrate of approximately 30% H2O2 was freshly diluted with water to about 10 mM and the concentration was accurately determined by using the molar extinction coefficient of 43 M-1cm-1 at the 240nm absorbance wavelength (Long et al., 1999a). From this intermediate standard, further dilutions in water or buffer (depending on which of the below-mentioned assays was used) were carried out to obtain the concentration or range of concentrations of H2O2 necessary for the experiment or standard calibration of assay (usually between 0 to 100 µM). 21 2.2.2. Preparation of human subjects Subjects were on self-selected diet with no special restrictions imposed, but with the following two exceptions. Subjects were not allowed to drink coffee or tea for at least 14 hours before the experiment and during the time of experiment (unless coffee is part of the experiment). Subjects were told not to overindulge in just one or a few particular types of food one day before and during the time of experiment (for example, subjects do not make fruits, vegetables, chocolates or alcoholic beverages as a quantitatively major part of their diet). Most importantly, subjects must be physically well. 2.2.3. Oxygen electrode assay This assay was largely based on the method described by Long et al. (1999b). A Hansatech oxygen electrode (Hansatech, UK) was used. Processing of signals from the electrode and recording of raw data were accomplished using a PowerLab® system and Chart™ Software, both from ADInstruments, New Zealand. The electrode was set up (with saturated aqueous KCl as the electrolyte) and stabilized for 30 min with 1.5 ml of deionized water at room temperature (25oC) in the reaction chamber. The chamber was then emptied and filled with 1.5 ml of urine sample. After a stable baseline was recorded on the chart, 100 µl of catalase (of type and source specified in 2.1.1) solution (10,000 U/ml in PBS buffer) was introduced to the chamber through the plunger capillary hole. The net deflection was recorded on the chart and the urinary H2O2 calculated. The electrode was calibrated for O2 evolution using freshly-prepared solutions of H2O2 in water (1.5 ml each) of known concentrations. 22 2.2.4. Recovery study for oxygen electrode assay Urine was freshly voided in 50-ml Greiner tubes from different individuals as well as the same individuals but on different days so that a total of 8 different samples was obtained. Only one urine sample was studied at a time. Since the concentration of each urine was different, the creatinine level was also determined where possible (refer to section 2.2.24). 1.5 ml of neat urine was introduced into the O2 electrode chamber and analyzed as described in 2.2.3. After rinsing the chamber clean, the experiment was repeated with more 1.5 ml portions of neat urine with one additional step: varying volumes of 5 mM H2O2 in water were added into the urine as well and dispersed by the magnetic stirrer. The following table showed the volume of 5 mM aqueous H2O2 added to 1.5 ml of urine to achieve the corresponding desired concentration of spiked H2O2. Conc. of spiked H2O2 (µM) Volume of 5 mM H2O2 (µl) neat 0.0 5 1.5 10 3.0 15 4.5 20 6.0 30 9.0 Each recovery was then calculated based on the response to the spiked H2O2 alone and not the total urinary H2O2 + spiked H2O2, i.e. % recovery = 100(A – B)/C where A = experimentally-determined total concentration of H2O2 in the spiked urine, B = experimentally-determined concentration of H2O2 in the neat urine and C = theoretical concentration of spiked H2O2 alone in the spiked urine. The neat urine was analyzed again at the end of each spiking experiment to check for any significant increase in the level of endogenous H2O2. 23 2.2.5. Study of ascorbate effect on oxygen electrode assay (a) Constant [ascorbate] and varying [spiked H2O2] Urine was freshly voided in a 50-ml Greiner tube from one individual and transferred into 2 separate tubes, so that each tube contained 20 ml of urine. 125 µ l of freshly-made 40 mM sodium L-ascorbate (Mr = 198.1) solution was added to one tube and 125 µl of water was added to the other. 1.5 ml of the 0.25 mM ascorbate-added urine was analyzed with the O2 electrode. Subsequently, 1.5 ml volumes of this urine but spiked with varying volumes of 5 mM H2O2 in water were analyzed as described in 2.2.4. The procedure was repeated with the control urine (without externally-added ascorbate). At the end, the recovery percentages of varying levels of spiked H2O2 were calculated for both the control and the 0.25 mM ascorbate-added urine. (b) Varying [ascorbate] and constant [spiked H2O2] Urine was freshly voided in a 50-ml Greiner tube from one individual and transferred separately into five 6-ml tubes. Different volumes of freshly-made 40 mM sodium L-ascorbate solution were added to the 5 tubes so as to achieve the following concentrations of ascorbate in urine: 0, 0.05, 0.10, 0.20 and 0.40 mM. Each 6-ml tube of urine was analyzed unspiked as well as spiked with an additional 10 µM of H2O2 (3 µl of 5 mM H2O2 in water was added to 1.5 ml of sample) using the O2 electrode. At the end, the recovery percentage of 10 µM spiked H2O2 for each unique concentration of ascorbate in urine was calculated like in 2.2.4. 24 2.2.6. Ferrous ion oxidation- xylenol orange version 2 (FOX-2) assay The FOX-2 assay (Long et al., 1999b and 2000) is based on the oxidation of Fe2+ by H2O2 to Fe (III), which then forms a measurable complex with xylenol orange. The following two reagents were prepared. Reagent 1 was 4.4 mM butylated hydroxytoluene (BHT) in methanol; reagent 2 was 1 mM xylenol orange plus 2.56 mM ferrous ammonium sulphate in 250 mM H2SO4. One volume of reagent 2 was mixed with nine volumes of reagent 1 to make the FOX-2 reagent which was stored in the dark at 040C for not more than a month. Urine sample or water (90 µl) was mixed with 10 µ l of methanol and vortexed. 900 µl of FOX-2 reagent was added, vortexed and incubated for 10 minutes at room temperature. Solutions were then centrifuged at 15000 g for 10 min at 4oC. The absorbance at 560 nm was read against a methanol blank. As controls, the above procedures were repeated with urine samples but adding 10 µl of catalase solution (1000 U/ml in PBS buffer) instead of methanol. The FOX-2 reagent was calibrated with known concentrations of hydrogen peroxide in water. Calculation of concentration of H2O2 in sample was done as the following:If Absλ=560nm (90µl water + 10µl MeOH + 900µ l FOX-2 reagent) = AWM, Absλ=560nm (90µl sample + 10µ l MeOH + 900µl FOX-2 reagent) = ASM, Absλ=560nm (90µl water + 10µl catalase + 900µl FOX-2 reagent) = AWC and Absλ=560nm (90µl sample + 10µ l catalase + 900µl FOX-2 reagent) = ASC, then Absλ=560nm due to H2O2 in sample = (ASM - AWM) - (ASC - AWC) = AT So, concentration of H2O2 in sample = AT / (gradient of calibration plot). 25 2.2.7. Recovery study for FOX-2 assay Urine was freshly voided in 50-ml Greiner tubes from different individuals as well as the same individuals but on different days so that a total of 10 different samples was obtained. Only one urine sample was studied at a time. Since the concentration of each urine was different, the creatinine level was also determined where possible (refer to section 2.2.24). The urine sample was then aliquoted equally into six 6-ml tubes. Varying volumes of 5 mM and 10 mM H2O2 in water were added to the tubes so as to achieve the following concentrations of spiked H2O2 in urine: 0, 5, 10, 20, 30 and 40 µM. These were then analyzed and calculated by similar procedures stated in 2.2.6, and the recovery percentages were calculated as described in 2.2.4. 2.2.8. FeTMPyP-catalysed indamine dye formation assay (FeTMPyP assay) This assay was a modification of the method described by Masuoka et al. (1996). It made use of an iron porphyrin to catalyse the H2O2-dependent formation of an indamine dye by oxidative coupling of N,N-dimethylaniline and a hydrazone (Fig. 3.5). (a) Initial Attempts A reagent solution was prepared by mixing equal volumes of the following: (1) 0.2 mM solution of FeTMPyPCl5 [meso-tetrakis(1-methyl-4pyridyl)porphinatoiron(III) pentachloride] (Mr = 909.9) in water, (2) 41.2 mM N,N-dimethylaniline (Mr = 121.18) in 0.2 M HCl and (3) 8.56 mM 3-methyl-2-benzothiazolinone hydrazone hydrochloride (Mr = 233.72) in 0.2 M HCl. The reagent solution was used within 3 hours after mixing. 26 500 µl of working standard or sample was mixed with 5 µ l of 0.2 M HCl and vortexed. 500 µl of reagent solution was added, vortexed and incubated for 1 hour at room temperature. Solutions were then centrifuged at 15000 g for 10 min at 4oC. The absorbance at 590 nm was read against a 0.2 M HCl blank. As controls, the above procedures were repeated with 500 µ l of urine samples but adding 5 µ l of catalase solution (10,000 U/ml in PBS buffer) instead of 0.2 M HCl. The reagent solution was calibrated with known concentrations of hydrogen peroxide in water. Calculation of concentration of H2O2 in sample S was done as the following:If Absλ=590nm (500µl S + 5 µl 0.2 M HCl + 500µl reagent solution) = ASH, Absλ=590nm (500µl S + 5 µl catalase + 500µ l reagent solution) = ASC, then Absλ=590nm due to H2O2 in sample S = ASH – ASC = AT So, concentration of H2O2 in sample S = AT / (gradient of calibration plot) (b) Use of higher reactant concentrations Basically, (b) differed from (a) in that the concentrations of FeTMPyP, 3-methyl2-benzothiazolinone hydrazone and N,N-dimethylaniline in the reaction mixture were increased several fold. The reagent solution was prepared by mixing the following in the stated volumes: (1) 1 volume of 0.4 mM solution of FeTMPyPCl5 in water, (2) 1 volume of 200 mM N,N-dimethylaniline in 0.2 M HCl and (3) 2 volumes of 25.7 mM 3-methyl-2-benzothiazolinone hydrazone hydrochloride in 0.2 M HCl. The remaining steps for samples and standards, and calculations were similar to that in (a). 27 2.2.9. Recovery study for FeTMPyP assay Urine was freshly voided in 50-ml Greiner tubes from the same individual but on different days so that a total of 3 different samples was obtained. Each day, the urine sample was aliquoted equally into six 6-ml tubes. Varying volumes of 5 mM H2O2 in water were added to the tubes so as to achieve the following concentrations of spiked H2O2 in urine: 0, 2.5, 5, 10, 20 and 40 µM. These were then analyzed and calculated using the same procedures given in 2.2.8. The recovery percentages were calculated as described in 2.2.4. 2.2.10. Pentafluorobenzenesulfonyl fluorescein (PFBSF) assay PFBSF was first designed and used for H2O2 analyses by Maeda et al. (2004). The assay is based on the H2O2-assisted cleavage of the pentafluorobenzene sulfonate moiety in PFBSF to release fluorescein (Fig. 3.9). PFBSF was dissolved in DMSO to give a 10 mM solution. This was diluted 100 times with cold (4oC) HEPES buffer (pH 7.4, 10 mM) to form the reagent solution (0.1 mM). The reagent solution was used as soon as it was prepared. The following two experiments were carried out:(1) 150 µl of reagent solution was mixed with 50 µl of H2O2 solution in HEPES buffer in a 96-well plate and incubated for 45 minutes at 37oC. The fluorescence was measured using λexcitation = 498 nm and λemission =522 nm. 0, 0.5, 1, 2, 3, 4, 5, 6 and 7 µM H2O2 standards were used. 28 (2) 110 µl of reagent solution was mixed with 90 µl of H2O2 solution in HEPES buffer in a 96-well plate and incubated for 45 minutes at 37oC. The fluorescence was measured using λexcitation = 498 nm and λemission =522 nm. 0, 10, 20, 30, 40, 50, 60, 70, 80 and 100 µM standards were used. 2.2.11. Homovanillic acid (HVA) assay This is a peroxidase-based assay using homovanillic acid (HVA; Mr = 182.18) as the oxidizable substrate (Fig. 3.10). HVA was dissolved in water to give a 10 mM solution. Horseradish peroxidase (HRP) was dissolved in PBS to give a 1000 U/ml solution. Appropriate volumes were mixed and diluted with more PBS to give a reagent solution of the following content: 0.3 mM HVA and 4.5 U/ml HRP. For each sample, 1 ml of (neat or diluted) sample was mixed with 10 µl of PBS and another 1 ml was treated with 10 µl of 2000 U/ml catalase. Both the reaction and catalase-treated mixtures were vortexed and incubated at room temperature for 1 min. 100 µl of reagent solution was mixed with 100 µl of treated sample or H2O2 standard in PBS on a 96-well plate and incubated for 2 min. The fluorescence of the mixture was measured at λexcitation = 312 nm and λemission = 420 nm (Barja, 2002). 2.2.12. p-Hydroxyphenyl acetic acid (HPAA) assay In this assay, p-hydroxyphenyl acetic acid (HPAA; Mr = 152.15) was used as the oxidizable substrate, instead of HVA. All steps were similar to that of the HVA assay (2.2.11) with the only exception being the final step where the fluorescence of the reaction mixture was measured at λexcitation = 317 nm and λemission = 414 nm. 29 2.2.13. 2,2’-Azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) assay [ABTS assay] Here, 2,2’-Azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) was used as the peroxidase substrate (Fig. 3.13). ABTS, diammonium salt (Mr = 548.68) and HRP were dissolved in PBS to give a reagent solution of the following content: 2 mM ABTS and 8 U/ml HRP (Li et al., 2002). 200 µl of working standard or urine sample was mixed with 10 µ l of PBS and vortexed. 790 µl of reagent solution was added, vortexed and incubated for 5 minutes at room temperature. The absorbance was read against a PBS blank at 730 nm (Yang et al., 2005). As controls, the above procedures were repeated with urine samples but adding 10 µ l of catalase solution (2,000 U/ml in PBS buffer) instead of PBS. The reagent solution was calibrated with known concentrations of hydrogen peroxide in water. 2.2.14. Preformation of ABTS+• and the quenching effect of urine The ABTS cation radical was preformed using the method described by Re et al. (1999). Firstly, the following two reagents were prepared. Reagent 1 was 7 mM ABTS in water. Reagent 2 was 24.5 mM potassium persulfate (K2S2O8) in water. One volume of reagent 2 was mixed with nine volumes of reagent 1 and the solution was left in the dark for 16 hours. ABTS and K2S2O8 react stoichiometrically at a ratio of 1:0.5, and required more than 6 hours to reach completion. The ABTS+• formed was stable for at least 2 days when stored in the dark at 40C. The ABTS+• solution was then diluted 50 times with PBS. 790 µl of the diluted ABTS+• solution was mixed with 200 µ l of PBS or urine sample (diluted 20 times with PBS). 10 µl of PBS was added to bring the total reaction 30 volume to 1 ml and the reaction mixture was incubated for 3 minutes at room temperature. The absorbance was read against a PBS blank at 730 nm. 2.2.15. N-Acetyl-3,7-dihydroxyphenoxazine (amplex red) assay As shown in Fig. 3.18, amplex red (Mr = 257.25) could be used as a peroxidase substrate to detect H2O2. Amplex red was dissolved in DMSO to give a 50 mM solution. HRP was dissolved in PBS to give a 1000 U/ml solution. Appropriate volumes were mixed and diluted with more PBS to give a reagent solution of the following content: 0.16 mM amplex red and 3 U/ml HRP. Any remaining unused 50 mM amplex red stock was pipetted into 0.6 ml-microfuge tubes and stored at – 20oC in the dark. For each sample, 1 ml of (neat or diluted) sample was mixed with 10 µl of PBS and another 1 ml was treated with 10 µl of 2000 U/ml catalase. Both the reaction and catalase-treated mixtures were vortexed and incubated at room temperature for 1 min. 150 µl of reagent solution was mixed with 50 µl of treated sample or H2O2 standard in PBS on a 96-well plate and incubated for 2 min. The fluorescence of the mixture was measured at λexcitation = 563 nm and λemission = 587 nm (Zhou et al., 1997). Calculation of concentration of H2O2 in sample S was done as the following:Concentration of H2O2 in sample S = 1.01ND (FS - FC) / m where FS = relative fluorescence units (RFU) of reaction mixture, FC = RFU of catalase-treated mixture, ND = no. of times dilution of urine sample, m = gradient of standard calibration plot, and 31 1.01 is a factor to take into account the dilution brought about through the introduction of 10 µl of PBS or catalase to the sample prior to reaction with reagent solution. 2.2.16. Recovery study for amplex red assay Urine was freshly voided in 50-ml Greiner tubes from different individuals as well as the same individuals but on different days so that a total of 9 different samples was obtained. Only one urine sample was studied at a time. Since the concentration of each urine was different, the creatinine level was also determined at the end (refer to section 2.2.24). The urine sample was aliquoted equally into six 6-ml tubes. Varying volumes of 5 mM and 10 mM H2O2 in water were added to the six tubes of neat urine. Portions of the spiked/unspiked urine were then diluted by a certain number of times approximated based on the observed intensity of colouration of the urine sample (i.e. the more concentrated the urine appeared, the higher the number of times of dilution), so that after dilution, the following concentrations of spiked H2O2 were achieved: 0, 0.5, 1, 2, 3 and 4 µM. The spiked and unspiked samples were then treated by the same procedure mentioned in 2.2.15 and the fluorescence measurements taken. Each recovery was then calculated based on the response to the spiked H2O2 alone, i.e. % recovery = 100 (A’ – B’)/C’ where A’ = experimentally determined total concentration of H2O2 in the diluted spiked urine, B’ = experimentally-determined concentration of H2O2 in the diluted unspiked urine, and C’ = theoretical concentration of spiked H2O2 in the diluted spiked urine. A’ and B’ can be individually calculated using the formula in 2.2.15 as a guide. 32 2.2.17. 2’,7’-Dichlorodihydrofluorescein (DCFH) assay 50 mg of 2’,7’-dichlorodihydrofluorescein diacetate (DCFH-DA; Mr = 487.3) was dissolved in DMSO to give a 50 mM stock solution. 50µ l portions were aliquoted into several 0.6 ml-microfuge tubes and stored at -20oC in the dark. Immediately before use, one tube of stock solution was thawed in the dark and then mixed with an equal volume of 0.1 M NaOH as well as 20 µl of ethanol. The mixture was left in the dark at room temperature for 30 minutes. The purpose of this step was to allow the hydrolysis of DCFH-DA to 2’,7’-dichlorodihydrofluorescein (DCFH) (Hempel et al., 1999). DCFH could then be used as a HRP substrate to measure H2O2 in this assay (Fig. 4.1). HRP was dissolved in PBS to give a 1000 U/ml solution. Appropriate volumes of freshly-formed DCFH and HRP solutions were mixed and diluted with more PBS to give a reagent solution of the following content: 0.16 mM DCFH and 3 U/ml HRP. For each sample, 1 ml of (neat or diluted) sample was mixed with 10 µl of PBS and another 1 ml was treated with 10 µl of 2000 U/ml catalase. Both the reaction and catalase-treated mixtures were vortexed and incubated at room temperature for 1 min. 150 µl of reagent solution was mixed with 50 µl of treated sample or H2O2 standard in PBS on a 96-well plate and incubated for 10 (or 30) minutes. The fluorescence of the mixture was measured at λexcitation = 498 nm and λemission = 522 nm (Gomes et al., 2005). Urinary H2O2 concentration was calculated using the formula given at the end of 2.2.15. 33 2.2.18. Recovery study for 2’,7’-dichlorodihydrofluorescein (DCFH) assay All the steps taken for this study and calculations were similar to that in 2.2.16, but with the following differences: (a) 14 different samples were collected. (b) The sample treatment, reaction and measurements were done according to that described in 2.2.17. 2.2.19. Monitoring the progress of DCFH assay and the effect of catalase and SOD (a) Reaction with working standards The following concentrations of H2O2 working standards were prepared in PBS: 0, 1, 2, 3, 4, 5, 6, 8 and 10 µM. 50 µl of each standard concentration was mixed with 150 µ l of DCFH reagent solution (as prepared in 2.2.17) on a 96-well plate and the reaction was monitored immediately for 13 minutes. For every 30 seconds, the fluorescence of the reaction mixture was measured at λexcitation = 498 nm and λemission = 522 nm. (b) Reaction with samples and study of effect of catalase and SOD Two urine samples (S1 and S2) from two individuals were freshly voided in 50ml Greiner tubes. Each sample was aliquoted into 6 microfuge tubes (labeled S1A to S1F and S2A to S2F) and diluted 2X to a total volume of 1 ml each. Catalase and SOD solution in PBS (2000 U/ml each) were prepared separately. Each tube of S1 and S2 would then receive the following additional treatments: (A) 20 µl PBS, (B) 20 µl SOD solution, (C)10 µl SOD & 10 µl PBS; (D) 20 µl catalase solution; (E) 10 µl catalase & 10 µ l PBS; (F) 10 µl catalase & 10 µ l SOD solutions. The tube mixtures were vortexed and incubated at room temperature for 1 min. 50 µl from each tube was mixed with 150 µl of 34 DCFH reagent solution (as prepared in 2.2.17) on a 96-well plate and the reaction was monitored immediately for 35 minutes. For every 5 minutes, the fluorescence of the reaction mixture was measured at λexcitation = 498 nm and λemission = 522 nm. 2.2.20. Dihydrorhodamine 123 (DHR) assay In the final assay, dihydrorhodamine 123 (DHR; Mr = 346.4) was used as the HRP substrate for H2O2 measurements. 10 mg of DHR was dissolved in DMSO to obtain a 25 mM stock solution. 50µ l portions were aliquoted into several 0.6 ml-microfuge tubes and stored at -20oC in the dark. HRP was dissolved in PBS to give a 1000 U/ml solution. Immediately before use, appropriate volumes of thawed stock DHR and HRP solution were mixed and diluted with 30% v/v methanol in PBS to give a reagent solution of the following content: 0.16 mM DHR and 3 U/ml HRP. For each sample, 1 ml of (neat or diluted) sample was mixed with 10 µl of PBS and another 1 ml was treated with 10 µl of 2000 U/ml catalase. Both the reaction and catalase-treated mixtures were vortexed and incubated at room temperature for 1 min. 150 µl of reagent solution was mixed with 50 µl of treated sample or H2O2 standard in PBS on a 96-well plate and incubated for 10 minutes. The fluorescence of the mixture was measured at λexcitation = 505 nm and λemission = 529 nm (Gomes et al., 2005). Urinary H2O2 concentration was calculated using the formula given at the end of 2.2.15. 35 2.2.21. Recovery study for dihydrorhodamine 123 (DHR) assay All the steps taken for this study and calculations were similar to that in 2.2.16, but with the following differences: (a) Only one sample was collected. (b) The sample treatment, reaction and measurements were done according to 2.2.20. (c) DCFH assay was also used on this sample for the purpose of comparison. 2.2.22. Basal urinary hydrogen peroxide measurements in human subjects Preparation and requirements of human subjects involved in this study are stated in 2.1.2 and 2.2.2. Urine samples were collected at 1100hrs and every two hours thereafter up till 1700hrs. Urine samples were freshly-voided in 50-ml Greiner tubes and the H2O2 concentrations were determined by the DCFH and O2 electrode assays as soon as possible. The H2O2 concentrations were normalized with creatinine concentration (refer to section 2.2.24). Experiments on each subject were repeated over 3 separate days within a period of 6 months. 2.2.23. Coffee drinking study One of the subjects involved in 2.2.22 was selected for this study. Urine sample was collected from him in a 50-ml Greiner tube at 1100hrs. Coffee (refer to section 2.1.3.) was consumed immediately after this collection and completely drunk within 5 minutes. Thereafter, urine samples were collected again at 1130, 1200, 1230, 1300, 1500 and 1700 hrs. Urinary H2O2 concentrations were determined by the DCFH and O2 electrode assays as soon as possible. The H2O2 concentrations were normalized with 36 creatinine concentration (refer to section 2.2.24). The experiment was repeated over 3 separate days within a period of 6 months. 2.2.24. Creatinine assay The method was described by Sigma Chemical Corp. The creatinine colour reagent was prepared by mixing the following components:(a) 3 volumes of stock picric acid (1% solution in water) (b) 1 volume of phosphate buffer (made up of 0.1 M Na2HPO4 and 0.1 M boric acid, pH adjusted to 7.4 with phosphoric acid) (c) 1 volume of 8% SDS The reaction was started by mixing the following in a 1.5-ml cuvette:(a) 75 µl of one of the following: (i) water (for blank), (ii) 3.0 mg/dl standard creatinine (for one-point standard), or (iii) urine sample that had been diluted by 20X. (b) 750 µl of creatinine colour reagent. (c) 150 µl of 1 M sodium hydroxide. The reaction mixture was incubated for 10 min. The absorbance at 500 nm was read against water-creatinine colour reagent blank. After reading, 25 µl of 60% acetic acid was added to each cuvette mixture and incubated for another 5 min before another set of absorbance readings were taken at 500 nm. The creatinine concentration in urine was then calculated using the following:Creatinine (mM) = [(initial sample absorbance – final sample absorbance) / (initial standard absorbance – final standard absorbance)] x 3 x 20 x 0.0884 37 CHAPTER 3 RESULTS AND DISCUSSION 3.1. CATALASE-BASED ELECTROCHEMICAL METHOD 3.1.1. Oxygen electrode assay B A Fig. 3.1. O2 electrode chart recording. An excerpt of a chart recorded for an experiment performed using the O2 electrode assay to illustrate key features. The upper half of the figure shows the deflections (for example from point A to B) where the y-axis represents percentage oxygen (% O2) and the x-axis represents the time scale in mins. Point A coincides with the addition of 1000 U of catalase into the solution in the chamber after attaining a stable baseline. An O2 burst results due to the catalytic decomposition of H2O2 in the solution, giving a deflection of magnitude proportional to the concentration of H2O2. When the decomposition completes, the baseline starts to stabilize again at Point B. The lower half of the figure shows the corresponding rate of change of percentage O2 with time (y-axis: %O2/s) and the time scale in mins as the x-axis. Three separate analyses of increasing H2O2 concentrations (neat urine, and urine spiked with 10 and 20 µM H2O2, respectively) are shown. 38 Reaction Upon the introduction of freshly voided urine into the chamber of the O2 electrode, the percentage oxygen level (% O2) fell rapidly and then stabilized. The decrease in % O2 is due to the hypoxic nature of urine when freshly-voided. When 1000 U of catalase was introduced into urine in the chamber, the stable baseline shifted upwards to give a deflection which later stabilized at a higher % O2. The deflection reflects a sudden burst of O2 production due to the catalytic decomposition of H2O2, based on the reaction: 2H2O2 → 2H2O + O2. The magnitude of the deflection was found to be proportional to the concentration of H2O2 in the chamber solution. Thus, urinary H2O2 concentration can be calculated after analyzing the magnitudes of baseline shifts for a range of standard H2O2 concentrations in water. Fig. 3.1 illustrates the chart recording of analyses of 3 different samples. Each sample decomposition reaction took not more than a minute while the entire analysis of each sample (including thorough rinsing of chamber before and after each analysis, equilibrating the sample in the chamber, reaction time and getting stable baselines) usually took about 5 minutes. Thus, for each freshly-voided urine sample, there was only sufficient time to do a duplicate analysis; anything more could affect the accuracy of the remaining samples waiting in the queue which have yet to be analyzed. Freshly-voided urine samples have to be analyzed immediately or as soon as possible because their values of H2O2 may change upon standing (Hiramoto et al., 2002 and Long et al., 1999b). 39 Standard Calibration Plot (O2 Electrode Assay) 12.0 Net Deflection 10.0 y = 0.1141x R2 = 0.9995 8.0 6.0 4.0 2.0 0.0 0 20 40 60 80 100 Concentration of H2O2 in µ M Fig. 3.2. A standard calibration plot for the O2 electrode assay. Typically 0, 10, 20, 30, 40 and 50 µM H2O2 solutions were used to calibrate for O2 evolution. In the above figure, however, three further concentrations were used to demonstrate the method’s linearity up to at least 80 µM while being able to attain an r2 (square of correlation coefficient) value of 0.999. Each data point is the mean ± SD of 3 separate experiments. Linearity of calibration plot 10, 20, 30, 40 and 50 µM of H2O2 in deionized water were routinely found to give measurable deflections that can be used to produce a standard calibration plot. Any deflections arising from water (0 µM of H2O2) were corrected to zero, and data from the other concentrations or samples were adjusted accordingly. In fact, the assay was found to be linear up to at least 80 µM of H2O2, as shown in Fig. 3.2. 40 Table 3.1. Accuracy of determination of PBS solutions of H2O2 by the O2 electrode assay. Two separate experiments (1 and 2) were carried out on different days. On both days, concentrations of 5 µM and below were difficult to determine accurately. Theoretical concentration of H2O2 in a PBS solution (µM) Experimentallydetermined H2O2 concentration in µM (accuracy in % is in parenthesis) 40 30 20 10 5 2.5 1 46.9 (117.2) 33.1 (110.3) 24.8 (124.1) 12.4 (124.1) 8.28 (165.5) 5.52 (220.7) 2 38.0 (94.9) 31.4 (104.7) 18.3 (91.6) 10.5 (104.7) 7.85 (157.1) 5.24 (209.4) Detection limit As shown in Table 3.1, there were no problems in accurately detecting H2O2 dissolved in PBS at concentrations between 10 to 40 µM. However, it was difficult to determine the 5 µM and 2.5 µM solutions due to the diminishing size of the deflections and sharpness of the peaks (in the %O2/s chart). The perceived H2O2 concentrations for both solutions were above 157% of the actual concentration. Thus, the O2 electrode assay is not sufficiently sensitive to measure urinary H2O2 at concentrations of 5 µM or less. Recovery data The neat and spiked urine samples were analyzed and the calculated recovery percentages are tabulated in Table 3.2. The eight different urine samples collected varied in their concentrations, as indicated by their creatinine content, ranging from a dilute one at 2.13 mM to a concentrated one at 18.1 mM. Nevertheless, all the urine samples consistently gave good recoveries of spiked H2O2, ranging from 75% to 120%. There was 41 no significant increase in the level of endogenous urinary H2O2 at the end of each spiking experiment. Thus, the recovery percentages were demonstrated to be independent of the urine concentration and the levels of spiking. Very importantly, the recovery studies showed that the O2 electrode assay is capable of reliably measuring urinary H2O2 accurately, and that the endogenous and spiked H2O2 is not lost by any reactions with the urinary constituents, i.e. urine does not appear to catabolize H2O2. Table 3.2. O2 electrode assay recovery study. Urine was freshly voided from different individuals as well as the same individuals but on different days so that a total of 8 different samples was obtained. For each urine sample collected, the neat form as well as those spiked with an additional 5, 10, 15, 20 and 30 µM H2O2 on top of the endogenous concentration was analyzed. The recovery percentages of spiked H2O2 were calculated as described in 2.2.4. Subject Sex M Urinary H2O2 (µM) 36.3 Urinary creatinine (mM) nd RD300905 Concentration of spiked H2O2 (µM) and their respective recoveries (%) 5 µM 10 µM 15 µM 20 µM 30 µM 80.6 80.6 116.5 107.5 98.6 RD061005 M 13.4 nd 89.4 96.8 89.4 93.9 100.6 RD281205 M 10.6 nd 105.5 105.5 112.1 109.9 108.8 MR171106 F 22.3 2.13 102.8 111.4 108.5 111.4 114.3 WH171106 M 40.3 15.3 85.7 94.3 102.8 111.4 111.4 NL171106 M 28.3 18.1 102.8 94.3 120.0 111.4 117.1 TS201106 M 15.8 4.96 81.7 76.6 74.9 84.3 78.3 RD201106 M 37.3 14.6 102.1 91.9 98.7 99.6 93.6 nd: not determined in experiment 42 Study of ascorbate effect on assay Many people take dietary supplements regularly or consume fruits and vegetables as part of their daily diet. Such foods and supplements can contain high amounts of ascorbic acid and excess ascorbate (AscH-) unutilized by the body is excreted in urine. Human urine was reported to contain between 0.15 to 0.18 mM of AscH- (Fang et al., 2006; Koshiishi et al., 2006). It will be interesting to examine the effect of AscH-, if any, on urinary H2O2 determination by the O2 electrode assay. Two approaches were taken in this study. In the first one, AscH- was added to urine and kept at a fixed concentration while H2O2 was added in varying amounts into the AscH--containing urine. In the second approach, H2O2 was spiked into urine at a fixed concentration while the amount of added AscH- was varied over a range of values. Both approaches would ultimately look at the percentage recoveries of spiked H2O2. Table 3.3. First study of ascorbate effect on O2 electrode assay. Urine was freshly voided from one individual and divided into two equal volumes. To one volume, ascorbate (AscH-) was added to give a concentration of 0.25 mM. AscH--containing urine was then analyzed unspiked as well as spiked with an additional 5, 10, 15, 20 and 30 µM of H2O2 on top of the endogenous concentration, using the O2 electrode. The procedure was repeated with the second portion (control urine, with no added AscH-). The recovery percentages of spiked H2O2 were calculated as described in 2.2.4. Sample 0.25 mM ascorbate added urine control urine Urinary H2O2 (µM) Urinary creatinine (mM) 18.2 6.69 91.2 100.3 109.4 109.4 118.5 23.6 6.69 118.2 109.8 107.0 114.0 95.7 Concentration of spiked H2O2 (µM) and their respective recoveries (%) 5 µM 10 µM 15 µM 20 µM 30 µM * At the end of the experiment, the urinary H2O2 of AscH--containing urine and control urine were measured again and found to be 21.1 and 22.8 µM respectively, while both their measured pHs were 6.0. 43 So, in the first study, one freshly-voided urine sample was split into two equal volumes, with one becoming the 0.25 mM AscH--added urine and the other as the control. Together, their recovery percentages of spiked H2O2 at five different levels ranged between 91.2% and 118.5% (Table 3.3). The AscH--added urine’s recovery percentage averaged 105.7 ± 10.4 % and the control urine’s recovery averaged 109.0 ± 8.5 % (mean ± SD, n=5), i.e. the recovery percentages of spiked H2O2 for both treatments were close to 100%. Very importantly, the addition of AscH- to urine does not alter the basal level of urinary H2O2; at the end of the first study, the urinary H2O2 of AscH-containing urine and control urine were found to be closely similar at 21.1 and 22.8 µM respectively, and the small difference between the basal readings of the two treatments during the spike recovery experiments (18.2 and 23.6 µM respectively) can be attributed to the production of more H2O2 in the collected urine sample upon standing, as the time interval between these 2 measurements were 30 minutes apart. The pH of the urine sample was 6.0, regardless of whether AscH- was present or not. In the second study, freshly-voided urine was split equally among five tubes and AscH- was added in the range of 0 to 0.40 mM [refer to 2.2.5(b) and Table 3.4]. The recovery percentages of 10 µM H2O2 spiked into these urine samples were in an acceptable range of 91.2% to 118.5% (Table 3.4). The data reflected no significant differences in H2O2 recovery between the different levels of added AscH- in urine. Added to that, the measured urinary H2O2 values before spiking (found to be in the range of 28.3 to 35.6 µM H2O2) were independent of the concentration of added AscH-. 44 Table 3.4. Second study of ascorbate effect on O2 electrode assay. Different amounts of ascorbate were added to separate portions of the same urine sample. Each of them was analyzed before and after the introduction of additional 10 µM H2O2 to the sample. Concentration of added ascorbate (mM) 0 0.05 0.10 0.20 0.40 Measured urinary H2O2 concentration (µM) After 10 µM H2O2 Before spiking spike 31.0 41.0 28.3 37.4 32.8 41.9 35.6 47.4 33.7 45.6 Recovery of 10 µM H2O2 spike (%) 100.3 91.2 91.2 118.5 118.5 * The pH of the urine sample is 5.8, regardless of the concentration of added ascorbate. The urinary creatinine concentration was 10.8 mM. O2 electrode assay: reliable, accurate but lacking sensitivity In short, the O2 electrode assay is simple, reliable and specific for measuring urinary H2O2, based on linearity of response and recovery data. It is not interfered by urinary ascorbate, up to a concentration of at least 0.40 mM, which is two times more than that usually detected in human urine. However, it lacks sensitivity and its accuracy is limited to H2O2 concentrations of above 5 µM. 45 3.2. NON-ENZYMATIC CHEMICAL-BASED METHODS 3.2.1. Ferrous ion oxidation- xylenol orange version 2 (FOX-2) assay Background Many groups used the ferrous ion oxidation- xylenol orange version 2 (FOX-2) assay to measure H2O2 in biological materials. After looking at how well the O2 electrode assay worked in measuring urinary H2O2, it was interesting to study the reliability of this commonly used assay for the same purpose. Reaction When FOX-2 reagent was added to a working standard of relatively high H2O2 concentration, a bluish violet colouration, observable to the naked eye, developed within the orange solution. This was due to H2O2 oxidising Fe2+ to Fe (III), which then formed a blue-violet chromogen with xylenol orange. Its absorbance maximum at 560nm can be used as a quantitative measure of the amount of H2O2 present. Linearity Fig. 3.3 shows a typical standard calibration plot for the assay. It was found to be linear in the range of 2.5 to 200 µM, with r2 > 0.99. Any absorbance at 560nm arising from water + FOX-2 reagent was corrected to zero and data from the other concentrations or samples were adjusted accordingly. 46 Standard Calibration Plot (FOX-2 Assay) 0.700 y = 0.0064x R2 = 0.9993 Net Absorbance 0.600 0.500 0.400 0.300 0.200 0.100 0.000 -0.100 0 20 40 60 80 100 120 Concentration of H2O2 in µ M Concentration of H2O2 in µM Total Absorbance Corrected Absorbance (560 nm) 0 0.096 0.000 (±0.012) 2.5 0.113 0.018 (±0.014) 5 0.129 0.034 (±0.012) 10 0.163 0.067 (±0.013) 20 0.226 0.130 (±0.018) Concentration of H2O2 in µM Total Absorbance Corrected Absorbance (560 nm) 30 0.294 0.199 (±0.019) 40 0.358 0.263 (±0.015) 50 0.421 0.325 (±0.017) 80 0.602 0.507 (±0.025) 100 0.723 0.627 (±0.027) Fig. 3.3. A standard calibration plot for the FOX-2 assay. In the above plot, 0, 2.5, 5, 10, 20, 30, 40, 50, 80 and 100 µM H2O2 in water were used to calibrate for Fe(III)-xylenol orange chromogen formation. Each data point is the mean ± SD of at least 3 separate experiments where in each experiment, duplicate measurements are made (the mean total absorbance and corrected absorbance at λ=560 nm as well as SD are tabulated below the plot). In fact, the relationship was found to be linear up to 200µM with an r2 > 0.99. Detection limit The FOX-2 assay is more sensitive than the O2 electrode assay as the former can detect as low as 2.5 µM of H2O2 while the detection limit of the latter is above 5 µM. 47 Recovery data The neat and spiked urine samples were analyzed and the calculated recovery percentages are tabulated in Table 3.5. Only 3 out of 10 samples have most of their recovery percentages of different levels of spiked H2O2 lying between 80 to 105% while another one had recoveries in the range of 70.0 to 81.2 %. The 3 worst samples have most of theirs ranging between 36.2 to 48.4%. In the remaining 3 samples, the recovery of spiked H2O2 ranged between 57.0 to 72.9%. Table 3.5. FOX-2 assay recovery study. Urine was freshly voided from different individuals as well as the same individuals but on different days to give a total of 10 different samples. The neat form of each urine sample, as well as those spiked with an additional 5, 10, 20, 30 and 40 µM H2O2 on top of the endogenous concentration, were analyzed. The recovery percentages of spiked H2O2 were calculated as described in 2.2.4. Subject Sex M Urinary H2O2 (µM) 14.0 Urinary creatinine (mM) nd RD290705 RD071205 M 8.39 7.67 151.3 104.8 87.5 81.0 nd SR231106 F 8.41 11.2 57.5 48.4 44.3 47.0 47.6 TS231106 M 4.90 1.89 86.3 83.8 83.6 83.8 83.7 TS251106 M 18.8 16.4 9.34 36.2 37.5 39.5 42.1 RD251106 M 19.2 20.5 16.4 37.9 42.9 48.1 47.6 NL281106 M 18.3 19.6 63.4 59.8 57.3 57.0 57.6 RD281106 M 16.4 3.08 70.0 78.5 76.5 73.4 81.2 MR051206 F 2.16 2.27 60.0 70.8 69.1 71.3 72.9 MR041206 F 1.18 2.43 58.4 61.6 67.7 70.1 71.9 Concentration of spiked H2O2 (µM) and their respective recoveries (%) 5 µM 10 µM 20 µM 30 µM 40 µM nd 93.5 75.7 80.5 80.3 nd: not determined in experiment 48 The ten different urine samples collected varied in their concentrations, as indicated by their creatinine content, ranging from a very dilute one at 1.89 mM to a highly concentrated one at 20.5 mM. The creatinine content was found to be an approximate indicator of the performance of H2O2 recovery experiments. It was noted that 2 of the 3 samples with the best recovery percentages of H2O2 had their creatinine concentrations at relatively low levels of 1.89 and 7.67 mM (for the third one, creatinine was not determined) while 2 of the 3 worst samples had very high creatinine levels (16.4 and 20.5 mM respectively, while the third one was lower at 11.2 mM). The subject with the second highest creatinine concentration (19.2 mM) had recoveries of H2O2 between 57.0 to 63.4% while the remaining three dilute samples (2.27, 2.43 and 3.08 mM of creatinine) had their values lying within a wide range of 58.4 to 81.2%. The recovery data of the FOX-2 assay was less impressive than that demonstrated by the O2 electrode assay. Some of the added H2O2 appeared to be ‘lost’ in the assay but a certain amount could still be detected. So, further investigations had to be carried out to find out the possible source of the problem. Comparison study between FOX-2 and O2 electrode assays The reliability of the O2 electrode assay was already observed earlier, so it was used in parallel with the FOX-2 assay in this study to analyze some urine samples. The time interval between the two assays was kept as short as possible, so as to reduce any discrepancies between them that might arise solely due to changes in endogenous urinary H2O2 with time, upon standing (Long et al., 1999b). 49 Table 3.6. Comparison of FOX-2 assay with O2 electrode assay in one individual. Urine samples were freshly voided from one individual at the times stated below on two different days and then analyzed for H2O2 using both the FOX-2 assay and the O2 electrode assay. FOX-2 assay data were derived from the average of duplicate absorbance measurements while O2 electrode assay data were the mean of 2 analyses. Date Time (hrs) 190605 060705 Urinary H2O2 in µM FOX-2 Assay O2 Electrode Assay 1000 8.62 10.5 (± 17.7%) 1100 5.13 16.0 (± 15.4%) 1200 5.49 12.9 (± 14.0%) 1300 12.8 17.2 (± 21.5%) 1400 25.4 30.8 (± 8.0%) 1500 13.2 33.2 (± 3.8%) 1600 38.5 22.1 (± 16.7%) 1400 11.7 31.1 (± 5.8%) 1500 12.8 34.8 (± 8.8%) 1600 13.4 46.4 (± 7.9%) 1700 16.4 44.5 (± 1.3%) 1800 8.86 33.6 (± 9.1%) The comparison study was done using two different approaches in order to study a variety of different urine samples. In the first one (Table 3.6), urine samples were freshly voided from one individual at regular time intervals on two different days. In the other approach (Table 3.7), spot urine samples were collected from various individuals only once within a day. Looking at Table 3.6, only three out of seven samples collected on 190605 (at 1000, 1300 and 1400 hours), had closely-agreeing values between the FOX-2 assay and 50 the O2 electrode assay. The four remaining samples collected on 190605 and all the five samples collected on 060705 gave very different values between the two assays. Table 3.7. Comparison of FOX-2 assay with O2 electrode assay in a few individuals. Urine samples were freshly voided from different individuals as well as the same individuals but on different days so that a total of 8 different samples was obtained. These were analyzed for H2O2 using both the FOX-2 assay and the O2 electrode assay. FOX-2 assay data were derived from the average of duplicate absorbance measurements while O2 electrode assay data were the mean of 2 analyses. Urinary H2O2 in µM FOX-2 Assay O2 Electrode Assay 8.41 20.7 (± 4.3%) Subject Sex SR231106 F TS231106 M 4.90 15.8 (± 2.9%) TS251106 M 18.8 25.7 (± 1.8%) RD251106 M 19.2 35.2 (± 2.6%) NL281106 M 18.3 21.9 (± 6.1%) RD281106 M 16.4 25.5 (± 1.8%) MR051206 F 2.16 9.66 (± 8.3%) MR041206 F 1.18 11.7 (± 3.4%) In Table 3.7, only three out of eight samples (TS251106, NL281106 and RD281106) had their measured urinary H2O2 values closely-agreeable between the two assays. One of the subjects had one sample (TS251106) where the FOX-2 and O2 electrode assays gave closely-agreeing values but another sample contributed two days earlier (TS231106) gave widely different values. Based on the latter finding and Table 3.6, it could be said that whether the two assays agreed or not did not depend on the identity of the individual but more likely on the content of various interfering compounds in the urine sample excreted at the particular time. 51 Combining the results of the two approaches together, only 30% of the total collected urine samples had closely-agreeing H2O2 values between the two assays. Very importantly, it was also noted that 19 out of the 20 samples had lower mean FOX-2 assay readings than those for the O2 electrode assay. Assuming that the O2 electrode assay is the true method for measuring urinary H2O2, the FOX-2 assay is constantly underestimating the actual H2O2 concentration in urine. This finding suggests the possible presence of substances in the urinary matrix that may have interfered with the actual determination of urinary H2O2 concentrations by the FOX-2 assay. Effect of dilution of urine samples on analyses If interferences arising from the urinary matrix were the cause of the frequent underestimations by the FOX-2 assay and the disagreements between the latter and the O2 electrode assay, it would be interesting to see if dilution of the samples could improve the accuracy of the FOX-2 assay by reducing the concentration of the interfering species in the reaction mixtures. The only difference here in terms of sample treatment compared to previous FOX-2 assay experiments was the dilution of the urine samples a specific number of times with water before 90 µl of the diluted form was aliquoted and further treated with the usual procedures before analyses. Two urine samples were collected from the same individual but on 2 different days, and another two urine samples were collected from 2 individuals on the same day. These samples were subjected to different numbers of dilutions as indicated in Table 3.8. The table shows the corresponding calculated concentrations of endogenous H2O2 in the undiluted sample for each level of dilution. 52 Table 3.8. Effect of dilution of urine sample on FOX-2 assay. Four urine samples were collected from 3 individuals. Each sample was subjected to the stated number of times of dilution with water before analyses and the corresponding calculated concentrations of endogenous H2O2 in the undiluted sample are given in the table. Subject RD011205 RD021205 TS020207 LH020207 Urinary creatinine (mM) 3.76 9.36 8.18 6.96 Number of times dilution and the calculated conc. of H2O2 in the undiluted sample (µM) Undiluted 2X 5X 10X 20X 31.8 33.3 30.6 28.7 29.3 5.65 9.78 13.4 21.2 18.3 17.2 27.7 32.5 30.4 22.2 6.37 8.27 11.3 8.33 11.7 Multiple dilutions of RD011205 made no difference to the calculated urinary H2O2 concentration and gave values similar to the undiluted sample. On the other hand, TS020207 and RD021205, which were 2.2 and 2.5 times more concentrated than RD011205, had higher and more consistent H2O2 values after 5 and 10 times dilution, respectively. LH020207 required only 2 times dilution. The data seems to suggest that dilutions of urine samples aid in the removal of significant amounts of interferences and thus enabling the detection of higher levels of urinary H2O2 by the FOX-2 assay. In order to confirm this, the recovery study, and the comparison study with the O2 electrode assay were repeated but with one major difference; samples were now diluted 10 times before FOX-2 assay was carried out (and so, will be denoted as 10xD-FOX-2 assay). However, neat urine samples were used for the O2 electrode assay. 53 Table 3.9. Comparison of 10xD-FOX-2 assay with O2 electrode assay. Urine samples were freshly voided from the same individual on six different days and analyzed for H2O2 using both 10xD-FOX-2 assay and O2 Electrode Assay. 10xD-FOX-2 assay data were derived from the average of duplicate absorbance measurements while O2 electrode assay data were given as mean ± SD of 3 independent measurements (with the exception of RD281205). Subject Urinary H2O2 in µM 10xD-FOX-2 Assay O2 Electrode Assay RD281205 13.3 10.6 RD271205 10.4 11.9 (± 0.8) RD151205 17.1 16.2 (± 1.3) RD141205 36.1 41.1(± 1.0) RD021205 21.2 14.3 (± 0.8) RD011205 28.7 12.6 (± 0.8) Unlike in Tables 3.6 and 3.7, where only 30% of the samples had closely-agreeing H2O2 values between the FOX-2 and O2 electrode assays, Table 3.9 showed that data from 10xD-FOX-2 assay tallied more closely with the O2 electrode data than that from FOX-2 assay. Five out of the six samples had comparable urinary H2O2 values between the two assays. The 10xD-FOX-2 assay values were no longer underestimates of the actual urinary H2O2 concentrations when compared with the O2 electrode assay. At the same time, the 10xD-FOX-2 assay also gave better recoveries of spiked H2O2 than the FOX-2 assay, ranging from 65% to 134% (Table 3.10). The most concentrated urine (RD091205 with 14.9 mM creatinine) had the lowest recovery percentages among the 3 samples, thus reinforcing the interfering role of some urinary compounds in H2O2 determinations by the FOX-2 assay. 54 Table 3.10. 10xD-FOX-2 assay recovery study. Urine was freshly voided from the same individual on 3 different days. For each sample collected, the neat form, as well as those spiked with an additional 5, 10, 15, 20, 50 and 100 µM on top of the endogenous concentration, was analyzed. The recovery percentages of spiked H2O2 were calculated as described in 2.2.4 Subject Sex M Urinary H2O2 (µM) 10.4 Urinary creatinine (mM) 3.69 RD271205 RD151205 M 17.1 6.43 nd 133.9 122.0 116.1 92.6 97.1 RD091205 M 62.7 14.9 65.6 64.8 79.2 113.5 83.8 nd Concentration of spiked H2O2 (µM) and their respective recoveries (%) 5 µM 10 µM 15 µM 20 µM 50 µM 100 µM 127.9 108.2 97.8 88.9 98.7 86.2 nd: not determined in experiment Problems with FOX-2 and 10xD-FOX-2 assays The data obtained from the recovery and assay comparison studies confirmed that dilutions of urine samples helped to diminish the effect of matrix interference by reducing its concentration in the reaction mixtures. Ascorbate is highly likely to be a main interfering species in the FOX-2 assay (Long, L.H., data not shown). Subjects who consumed more vegetables and fruits would have higher excretions of unutilized ascorbate in the urine. Ascorbate is known to be able to reduce Fe(III) to Fe2+ (Halliwell, 1996). The yield of Fe(III)-xylenol orange chromogen will actually be less than what the actual urinary H2O2 could have theoretically produced due to the reversion of some Fe(III) back to Fe2+, thus resulting in significant underestimation of H2O2 in such samples. The amount of interference was roughly correlated with the urine (creatinine) concentration. However, tweaking the FOX-2 assay by introducing sample dilution does not solve the problem with analyzing some urine samples. The detection limit of the FOX-2 55 assay is 2.5 µM. This means that only urine samples with a minimum of 25 µM H2O2 could be diluted 10 times and analyzed accurately by 10xD-FOX-2 assay. Many urine samples analyzed by the O2 electrode have been shown to contain less than 25 µM H2O2. But analyzing a range of dilution levels (from 2 to 10 times) for each urine sample can be too time-consuming, especially when many samples have to be analyzed. Increased preparation and analysis time per sample will increase the time lag between voiding of urine samples and actual analyses, such that the accuracy of pending sample measurements becomes questionable. By the same reasoning, H2O2 determination by the method of standard addition is also not a practical solution. Thus, the frequent underestimation of urinary H2O2 by the FOX-2 assay and the extremely low sensitivity of the 10xD-FOX-2 assay made both assays unsuitable for urinary H2O2 measurements. 56 3.2.2. FeTMPyP-catalysed indamine dye formation assay (FeTMPyP assay) Background Ferriprotoporphyrin IX (ferric heme or hemin; Fig. 3.4a) is the prosthetic group of most peroxidases and cytochrome P450, which by itself can react with H2O2 but has low solubility in water (Dunford, 1987). Nakano et al. (1990) created meso-tetrakis(1-methyl-4-pyridyl)porphinatoiron(III) complex (FeTMPyP; Fig. 3.4b), an artificial iron porphyrin which was stabilized by electron-withdrawing halogens like chlorine, and was water-soluble. The author used it to specifically catalyze the formation of an indamine dye by oxidative coupling of N,Ndimethylaniline and 3-methyl-2-benzothiazolinone hydrazone with H2O2 in an acidic media without any degradation of the hemin. The reaction equation is shown in Fig. 3.5. 57 Fig. 3.5. Coupling reaction to form indamine dye. Since the FOX-2 (and 10xD-FOX-2) assay was analytically validated with little success, it would be interesting to look at the above reaction as an alternative nonenzymatic, chemical-based method with a different mechanism of action to measure urinary H2O2. Fig. 3.6. Absorbance progress of the FeTMPyP-catalyzed indamine dye formation reaction. The reaction mixture was prepared as described in 2.2.8. The initial concentration of H2O2 was 70 µM. 58 Reaction Masuoka et al. (1996) allowed 1 hour of incubation for complete indamine dye formation by H2O2; this could not be further shortened as our investigation on the time taken for the complete reaction with 70 µM H2O2 took almost that amount of time (Fig.3.6). Reducing the sample or standard volume in the reaction mixture did not alter the reaction time (data not shown). Linearity Fig. 3.7 shows a typical standard calibration plot for the assay. It was found to be linear in the range of 2.5 to 60 µM, with r2 > 0.99. Any absorbance at 590nm arising from water + reagent solution was corrected to zero and data from the other working standards of H2O2 were adjusted accordingly. Concentrations of 70 µM and above started to curve out of the linear range (data not shown). Detection limit Like the FOX-2 assay, its detection limit of H2O2 is 2.5 µM. As can be seen in the table of data given together with Figure 3.7, the total absorbance (before correction) of 2.5 µM H2O2 reaction mixture (at 0.228 ± 0.025) was also very close to that of the control (water) reaction mixture (at 0.185 ± 0.020), so that if lower concentrations were analyzed, it would have been difficult to tell whether a net absorbance detected was due to real H2O2 present or just the result of random errors and fluctuations. 59 Standard Calibration Plot (FeTMPyP Assay) 0.900 y = 0.0139x Net Absorbance 0.800 R2 = 0.997 0.700 0.600 0.500 0.400 0.300 0.200 0.100 0.000 -0.100 0 10 20 30 40 50 60 70 Concentration of H2O2 in µ M Concentration of H2O2 in µM 0 2.5 5 10 20 Total Absorbance 0.185 0.228 0.269 0.342 0.486 Corrected Absorbance 0.000 0.043 0.084 0.156 0.300 (590 nm) (±0.020) (±0.025) (±0.029) (±0.027) (±0.029) Concentration of H2O2 in µM 30 40 50 60 Total Absorbance 0.618 0.751 0.870 0.995 Corrected Absorbance 0.433 0.566 0.685 0.809 (590 nm) (±0.032) (±0.036) (±0.040) (±0.041) Fig. 3.7. A standard calibration plot for the FeTMPyP assay. In the above plot, 0, 2.5, 5, 10, 20, 30, 40, 50 and 60 µM H2O2 in water were used to calibrate for indamine dye formation. Each data point on the linear plot was the mean ± SD of 3 separate experiments where in each experiment, duplicate measurements are made (the mean total absorbance and corrected absorbance at λ=590 nm as well as SD are tabulated below the plot). The relationship was found to be linear up to 60µM with an r2 > 0.99. Samples and recovery data Although the FeTMPyP assay worked very well for aqueous H2O2 as can be observed from the standard calibration plot, it did not work the same way for the samples. Endogenous H2O2 in neat urine samples was either detected at very low level or not detected at all (Table 3.11, samples RD150506 and RD110506). In fact, the absorbances 60 at 590 nm of ‘urine sample + reagent solution’ for both samples were lower than that of ‘water + reagent solution’ after 60 minutes of reaction, indicating that some other reactions might have possibly taken place between the urinary matrix and FeTMPyP. Recoveries of spiked H2O2 were dismal (in Table 3.11, between negative values to a maximum of 5.07% for samples RD150506 and RD110506). With the presence of an inherent source of interference in urine samples so major that almost all H2O2 in urine appeared quenched, the FeTMPyP assay did not seem suitable for urinary H2O2 analyses. Table 3.11. FeTMPyP assay recovery study. Urine was freshly voided from the same individual on different days to get a total of 3 different samples. For each urine sample collected, the neat form as well as those spiked with an additional 2.5, 5, 10, 20 and 40 µM H2O2 on top of the endogenous concentration was analyzed. The recovery percentages of spiked H2O2 were calculated as described in 2.2.4. Subject Urinary H2O2 (µM) Concentration of spiked H2O2 (µM) and their respective recoveries (%) 2.5 µM 5 µM 10 µM 20 µM 40 µM 5.97 7.69 3.92 0.43 1.70 RD080806 Not detected RD150506 0.15 5.07 4.48 0.15 0.07 0.06 RD110506 Not detected negative negative 2.36 1.37 1.37 Effect of use of higher reactant concentrations A large sample volume (500 µl) comprising half of the reaction mixture volume was required to attain the sensitivity and detection limit of 2.5 µM in the FeTMPyP assay. It is already seen with the FOX-2 assay that dilution of urine samples, while being able to reduce the concentration of interfering species, is going to unfavourably increase the detection limit in the same way and result in inaccurate determinations of low urinary H2O2 concentrations. Thus, another possible way to increase the probability of reaction of 61 urinary H2O2 with the reactants is to increase the reactant concentrations while keeping the volumes of the sample and total reaction mixture constant. The concentrations of FeTMPyP, 3-methyl-2-benzothiazolinone hydrazone and N,N-dimethylaniline in the reaction mixture were increased by 1.5, 4.5 and 3.6 times respectively. The absorbance at 590 nm of ‘water + reagent solution’ became higher due to the increased absorbance of the higher reactant concentration. But when this was corrected to zero with the other standards adjusted accordingly, the gradient of the calibration plot (y = 0.0159x) was close to and within 14% of that in Figure 3.7. Still, urinary H2O2 was not measurable (with its ‘reaction mixture’ absorbance still lower than ‘water + reagent solution’) and the recovery of spiked H2O2 ranged between 0.43 and 7.69 % (Table 3.11, sample RD080806). FeTMPyP assay: urinary matrix interference and problems The inability of the FeTMPyP assay to realistically detect endogenous and spiked H2O2 in human urine samples, even after increasing reactant concentrations by several fold, indicates the presence of a very significant interference arising from component(s) in the urinary matrix. Two possible constituents that could have contributed to the failure of the assay are urate and ascorbate. The initial reaction of Fe(III)TMPyP with H2O2 generates (TMPyP)•+Fe(IV)=O as a precursor of TMPyPFe(IV)=O (Saha et al., 2003a & b). The oxo-iron(IV) porphyrin generated could then participate in the oxidative coupling reaction between N,Ndimethylaniline and 3-methyl-2-benzothiazolinone hydrazone. Uric acid was found to be a scavenger of both (TMPyP)•+Fe(IV)=O and TMPyPFe(IV)=O (Saha et al., 2003a & b). 62 Added to that, the reduction of the (TMPyP)•+Fe(IV)=O back to Fe(III)TMPyP was more rapid than the formation of (TMPyP)•+Fe(IV)=O from Fe(III)TMPyP (k5 » k1) as well as the reduction of TMPyPFe(IV)=O to Fe(III)TMPyP (k5 » k6) in the presence of uric acid (Saha et al., 2003a & b) (Fig. 3.8). Fig. 3.8. FeTMPyP reaction scheme adapted from Saha et al. (2003a). k6 = 5.45 x 106 /mol/L/s and k1 =2.07 x 104 /mol/L/s. UA: uric acid. Considering that the oxidation of uric acid to allantoin by (TMPyP)•+Fe(IV)=O is simpler (involving only one molecule) and thus likely to be more rapid than the oxidative coupling of two molecules by TMPyPFe(IV)=O to form the indamine dye, the reaction depicted in Fig. 3.5 is not likely to proceed, at least not until complete oxidation of uric acid. Since the stoichiometry was found to be 1:1 for H2O2 and uric acid, indamine dye formation was not likely to be observed if the initial concentration of uric acid was the same or even more than that of H2O2. Human urine could contain between 0.4 to 4.4 mM of uric acid (Fang et al., 2006; Kalimuthu et al., 2006; Safavi et al., 2006), presumably depending on the level of hydration of the participant individuals. Since urinary H2O2 concentration would always be less than uric acid concentration, the FeTMPyP assay might not be predicted to detect any H2O2 in urine samples, regardless of the length of incubation time. 63 In the FOX-2 assay, ascorbate reduced Fe (III) to Fe2+ to prevent the formation of the Fe(III)-xylenol orange chromogen. Similarly, in the FeTMPyP assay, ascorbate could reduce the oxidation state of iron from +4 of TMPyPFe(IV)=O to +3 of Fe(III)TMPyP (Dunne et. al., 2006; Jensen et al., 2002), thus preventing the catalyst from carrying out its key role in the indamine dye formation reaction. It is not known whether ascorbate or urate played a greater role in the inhibition of the assay. 64 3.2.3. Pentafluorobenzenesulfonyl fluorescein (PFBSF) assay Background Acid form, X = H Lactone form Fig. 3.9. Pentafluorobenzenesulfonyl fluorescein (PFBSF). Compound is shown in acid and lactone forms, both existing in equilibrium in aqueous solution, and its perhydrolysis reaction with H2O2 (partially adapted from Maeda et al., 2004) The use of redox-based mechanisms to measure urinary H2O2 and the presence of chemical species in urine that directly interfered with these mechanisms led to the failure of the FOX-2 and FeTMPyP assays. Thus, the use of a probe for H2O2 that is based on a non-oxidative fluorescence mechanism in the next study seemed logical. Pentafluorobenzenesulfonyl fluorescein (PFBSF), which was synthesized by Maeda et al. (2004), had the potential to allow highly specific, peroxidase-independent detection of H2O2 under the complicated oxidative circumstances found in biological systems. It is based on a simple deprotection mechanism instead of oxidation. Fig. 3.9 illustrates the 65 structure of PFBSF and the perhydrolysis reaction. Sulfonates are more stable than esters to hydrolysis, with the pentafluorobenzene ring further enhancing the reactivity of the sulfonates toward H2O2 and not water (Maeda et al., 2004). PFBSF assay: no observable response to H2O2 standards However, a standard calibration plot could not be established with the PFBSF assay, at least for H2O2 concentrations of 100 µM and below. No improvements were observed when the sample volume was increased from 25% to 45% of the total reaction mixture volume in an attempt to increase the assay’s sensitivity. The background fluorescence of the reagent solution was already high to start with. It was observed that when the colourless DMSO solution of PFBSF (10 mM) was plunged into HEPES buffer to make the reagent solution (0.1 mM PFBSF), the solution turned yellow immediately without the introduction of H2O2. Although PFBSF was designed with the hope of at least significantly reducing the competition between perhydrolysis and hydrolysis, it was possible that significant hydrolysis of the compound still took place when making up the reagent solution, thus limiting or blocking the availability of intact PFBSF for perhydrolysis reaction with H2O2. Further precautions such as using cold (4oC) HEPES buffer to make the reagent solution, using the reagent solution immediately after preparation and limiting its exposure to light did not help. Still, there were no detectable changes in fluorescence of the reaction mixtures containing H2O2 working standards of concentrations between 0 to 100 µM, even after incubation for 45 min at 37oC. Either the increase in fluorescence intensity for each working standard was too small compared to the larger background fluorescence of the blank to be 66 detected, or H2O2 did not get to react with PFBSF. If the former is true, the detection limit of the PFBSF assay is far too high for the purposes of urinary H2O2 investigations. The difficulty of handling the probe due to its instability and susceptibility to hydrolysis made it not worthwhile to explore the assay any further. 67 3.3. PEROXIDASE-BASED METHODS None of the non-enzymatic chemical-based methods (FOX-2, FeTMPyP and PFBSF assays) were validated to be ideal for urinary H2O2 measurements. In this section of studies, peroxidase-based methods were explored. Peroxidase enzymes act on H2O2 by using it to oxidize another substrate (written as SH2 below) SH2 + H2O2 → S + 2 H2O SH2 are the non-fluorescent (or non-absorbing) probes used in the following studies to detect H2O2, which upon oxidation to S, will fluoresce at certain excitation/emission wavelengths (or absorb at a fixed wavelength), thus enabling the accurate quantitation of H2O2 at low µM levels. Peroxidase-based methods in various types of work are known for their high sensitivity. 68 3.3.1. Homovanillic acid (HVA) assay Background Homovanillic acid (4-hydroxy-3-methoxy-phenylacetic acid; HVA) is a nonfluorescent molecule that by reaction with H2O2, in the presence of horseradish peroxidase (HRP), produces a fluorescent dimer (Fig. 3.10). HVA is widely used for the detection and imaging of oxidative enzymes such as peroxidase (Foppoli et al., 2000), the determination of H2O2 scavenging activity (Pazdzioch-Czochra et al., 2002) and mitochondrial H2O2 generation (Barja, 2002). Reaction It took not more than a minute for the dimerization of HVA by H2O2 standards in the presence of horseradish peroxidase (HRP) to reach completion. Fig. 3.10. Oxidation of HVA in the presence of HRP to a fluorescence dimer (adapted from Pazdzioch-Czochra et al., 2002). HPAA, which differs structurally from HVA by the absence of the methoxy group at the meta position, participates in the same reaction. λexc/λemis for HPAA dimerization = 317/414nm. 69 Net Fluorescence Intensity Standard Calibration Plot (HVA Assay) 600 y = 49.067x R2 = 0.9991 500 400 300 200 100 0 0 1 2 3 4 5 6 7 8 9 10 11 Concentration of H2O2 in µ M Fig. 3.11. A standard calibration plot for the HVA assay. In the above plot, 0, 0.5, 1, 2, 3, 4, 5, 6, 8 and 10 µM H2O2 solutions in PBS were used to calibrate for HVA-dimer formation. Each data point is the mean ± SD of 3 separate experiments where in each experiment, duplicate measurements are made. The relationship was found to be linear up to at least 10 µM with an r2 > 0.99. Linearity Fig. 3.11 shows a typical standard calibration plot for the assay. It was found to be linear in the range of 0.5 to 10 µM, with r2 > 0.99. Any fluorescence arising from PBS + reagent solution was corrected to zero and data from the other standard concentrations were adjusted accordingly. Detection limit Finally, there was an assay that could detect as low as 0.5 µM of H2O2. 70 Table 3.12. Effect of dilution of urine sample on HVA assay. Two urine samples were collected from the same individual. Each sample was subjected to the stated number of times of dilution. Fluorescence intensities of reaction and catalase-treated control mixtures were found to be so similar that the endogenous H2O2 concentrations could not be reliably calculated. Each intensity value given is the average of two independent readings. No. of times dilution of urine 1 2 5 10 20 40 50 100 Relative Fluorescence Intensity (λexc/λemis =312/420 nm) Sample 011106a Sample 011106b Reaction Control Reaction Control 2551 2516 2200 2195 2168 2182 1694 1685 1469 1524 1034 1036 986 989 639 626 587 608 365 363 320 360 212 211 283 293 178 180 145 148 94.8 96.3 Samples data: problems with the HVA assay The HVA assay was better than the O2 electrode and all non-enzymatic chemicalbased assays discussed earlier by being able to detect H2O2 concentrations as low as 0.5 µM. Although the assay was very sensitive and linear in response to H2O2 in PBS as can be observed from its standard calibration plot, the assay did not work the same way with urine samples. Less than 1 µM of H2O2 was detected in two neat urine samples. Dilutions of urine samples between 2 to 100 times were carried out to reduce the concentration of any interfering species that might be present; even then, no H2O2 could be reliably detected, as the fluorescence emitted from both the reaction and catalasetreated mixtures were approximately similar (Table 3.12). Thus, the possibility that the urinary matrix blocks the formation of the HVA-dimer product cannot be dismissed. 71 When the reagent solution was substituted with PBS in the reaction mixture containing urine, the intensity of the fluorescence emission did not change significantly, thus indicating that most of the fluorescence was coming from the urinary matrix (Table 3.13). Furthermore, the fluorescence emitted from the two combinations containing neat urine was about 20 times higher than that from PBS + reagent solution, and 4 times higher than that from the highest standard employed (10 µM). This means that if there is any fluorescence emission from HVA dimerisation due to reaction with HRP and low amounts of urinary H2O2, it could have been easily eclipsed by the high fluorescence of the urinary matrix at the employed excitation/emission wavelengths. Attempts to improve sensitivity by using larger reaction volumes in 3-ml cuvettes (instead of 200 µ l) also failed as most of the detected fluorescence came from the urinary matrix itself (data not shown). Table 3.13. HVA assay: fluorescence in different mixtures. In every combination, 100 µl of each component was mixed (total volume = 200 µl). Each fluorescence intensity value is the average of two independent readings. Combination Urine + reagent solution Urine + PBS 10 µM H2O2 in PBS + reagent solution PBS + reagent solution Relative Fluorescence Intensity Reaction Catalase-treated Net mixture Mixture Difference 2180 (± 1.4%) 2200 (± 0.9%) 2105 (± 3.2%) 2169 (± 2.2%) 554.6 (± 1.1%) 90.0 (± 6.4%) 463.7 107.4 (± 4.2%) ≈ as above - Calculated [H2O2] / µM 11.1 - 72 3.3.2. p-Hydroxyphenyl acetic acid (HPAA) assay Background p-Hydroxyphenylacetic acid (HPAA) only differs structurally from HVA by the absence of the methoxy group at the meta position on the benzene ring. Like HVA, HPAA may turn out to be unsuitable for urinary H2O2 analyses. However, the difference in functional groups between the two compounds may cause them to interact differently with HRP intermediates and also influence the ease of the dimerization process (Fig. 3.10). This, together with the known sensitivity of peroxidase-based assays, makes the HPAA worth trying as a probe for urinary H2O2. Reaction, linearity and detection limit Like in the HVA assay, it took not more than a minute for the dimerization reaction to complete. The detection limit at 1 µM of H2O2 is slightly higher than that for the HVA assay (0.5 µM). But it still kept a linear relationship up to at least 10 µM H2O2, with r2 > 0.99 (Fig. 3.12). Similar problems Like in the HVA assay, it could not detect urinary H2O2. No H2O2 could be detected reliably with any number of dilutions between 2 to 100 times, as the fluorescence emitted from both the reaction and catalase-treated mixtures was not only approximately similar, but also very high when compared with that of the mixture of PBS + reagent solution (Table 3.14). 73 Standard Calibration Plot (HPAA Assay) Net Fluorescence Intensity 800 y = 71.923x R2 = 0.9999 700 600 500 400 300 200 100 0 0 2 4 6 8 10 12 Concentration of H2O2 in µ M Fig. 3.12. A standard calibration plot for the HPAA assay. In the above plot, 0, 1, 2, 3, 4, 5, 6, 8 and 10 µM H2O2 solutions were used to calibrate for HPAA-dimer formation. Each data point is the mean ± SD of 3 separate experiments where in each experiment, duplicate measurements are made. The relationship was found to be linear up to at least 10 µM with an r2 > 0.99. Table 3.14. Effect of dilution of urine sample on HPAA assay. Two urine samples were collected from the same individual on two different days. Each sample was subjected to the stated number of times of dilution. Fluorescence intensities of reaction and catalasetreated control mixtures were found to be so similar that the endogenous H2O2 concentrations could not be reliably calculated. Each intensity value given is the average of two independent readings. No. of times dilution of urine 1 2 5 10 20 40 50 100 Relative Fluorescence Intensity (λexc/λemis =317/414 nm) Sample 101106 Sample 091106 Reaction Control Reaction Control 3542 3581 4916 4892 2447 2427 3956 3946 1427 1473 2463 2527 897 901 1689 1692 675 623 1169 1165 434 450 784 755 348 380 621 660 244 294 476 475 74 It is perhaps not surprising to encounter similar problems when HPAA was used as a probe instead of HVA. Even if the difference in functional group substitution improved the kinetics of the reaction, the products of the dimerization process might still be detected with difficulty. After all, the excitation/emission wavelengths employed did not differ much and are found in the region where one experiences a lot of interferences from the urinary matrices. Thus, HVA and HPAA cannot be used as probes for urinary H2O2 analyses due to their low sensitivity of analyte detection coupled with high fluorescence emission from the sample matrix at the employed wavelengths. 75 3.3.3. 2,2’-Azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) assay [ABTS assay] Background A third peroxidase substrate, 2,2’-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) or ABTS, for short, was studied next. The ABTS assay involved the direct one-step oxidation of colourless ABTS by H2O2, in the presence of HRP, to form the blue-green ABTS monocation radical (ABTS+•) chromophore (Fig. 3.13). Fig. 3.13. Structure of ABTS and its oxidation products. In the presence of H2O2 and HRP, ABTS (I) can undergo a one-electron oxidation to give the metastable radical cation (II) or ABTS+•, which slowly disproportionates giving (I) and the azodication (III). This last chemical change is too slow to affect initial-rate measurements. (Adapted from Childs et al., 1975) 76 The λmax of ABTS+• (414 nm) lies at the region of visible spectrum that gives a lot of sample interference problems during the investigation of the HVA and HPAA assays, so the next best absorbance wavelength at 730 nm (ε725 = 14 200 M-1cm-1, Yang et al., 2005) was chosen. Reaction When 100 µM H2O2 was reacted in the ABTS/HRP system, the ABTS+• formed was stable for at least 60 min; its absorbance changed very little, from 0.568 to 0.559. Linearity Fig. 3.14 shows a typical standard calibration plot for the assay. It was found to have excellent linearity in the range of 2.5 to 200 µM, with r2 > 0.99. Any absorbance at 730 nm arising from water + ABTS reagent solution was corrected to zero and data from the other concentrations or samples were adjusted accordingly. Detection limit Like in FOX-2 and FeTMPyP assays, the ABTS assay was only able to accurately quantitate a sample containing at least 2.5 µM of H2O2. As can be seen in the table of data given together with Fig. 3.14, the total absorbance (before correction) of 2.5 µM H2O2 reaction mixture (at 0.023 ± 0.0%) was very close to that of the control (water) reaction mixture (at 0.013 ± 25.1%), so that if lower concentrations were analyzed, it would have been difficult to tell whether a net absorbance detected was due to real H2O2 present or just the result of random fluctuations. 77 Standard Calibration Plot (ABTS Assay) 1.2 y = 0.0056x R2 = 1 Net Absorbance 1 0.8 0.6 0.4 0.2 0 0 50 100 150 200 250 Concentration of H2O2 in µ M Concentration of H2O2 in µM Relative Absorbance (730 nm) 0 2.5 5 10 20 0.013 0.023 0.041 0.069 0.125 (±25.1%) (±0.0%) (±5.9%) (±6.9%) (±3.3%) Concentration of H2O2 in µM Relative Absorbance (730 nm) 30 40 50 100 200 0.180 0.237 0.289 0.568 1.12 (±3.1%) (±2.5%) (±3.3%) (±1.0%) (±1.4%) Fig. 3.14. A standard calibration plot for the ABTS assay. In the above plot, 0, 2.5, 5, 10, 20, 30, 40, 50, 100 and 200 µM H2O2 solutions were used to calibrate for ABTS radical monocation formation. Each data point is the mean of two separate experiments. The relative absorbance (before correction with respect to 0 µM H2O2) at λ=730 nm as well as % variation are tabulated below the plot. In fact, the relationship was found to be linear up to 200µM with an r2 > 0.99. Samples and recovery data The ABTS assay could not measure urinary H2O2 concentration at all. The absorbances of the reaction mixtures containing urine (spiked and unspiked) were far lower than those of the working standards and even the PBS mixture. Two urine samples studied gave similar observations, and Table 3.15 shows one of the samples’ data. 78 Recoveries of spiked H2O2 could not be calculated at all as the absorbances for both the reaction mixtures and catalase-treated controls were, at the same time, too low and similar to each other. Again, the failure of the assay is attributed to the composition of urine. Table 3.15. ABTS assay: sample absorbance data. One urine sample was freshly voided; the neat form as well as those spiked with an additional 5, 10, 20, 40 and 80 µM H2O2 on top of the endogenous concentration was analyzed. Relative absorbances of neat and spiked urine mixtures were tabulated below and found to be lower than those of the standards and even PBS blank. Each absorbance value is the mean of a duplicate analysis (± % variation). Reaction mixture Control mixture Neat PBS urine 0.0093 0.0020 (± 3.2%) (± 12.8%) 0.0089 0.0016 (± 11.2%) (± 3.2%) + 5 µM 0.0034 (± 2.9%) 0.0017 (± 3.0%) +10 µM + 20 µM + 40 µM + 80 µM 0.0043 0.0023 0.0033 0.0020 (± 44.2%) (± 21.7%) (± 29.2%) (± 5.0%) 0.0026 0.0020 0.0028 0.0026 (± 19.2%) (± 2.6%) (± 27.3%) (± 11.5%) Preformation of ABTS+• and the quenching effect of urine A question to be addressed was whether the end-product ABTS+• could be quenched in urine. It is known that ABTS+• is reduced in the presence of hydrogendonating antioxidants (Re et al., 1999). In fact, ABTS+• was the species generated by the reaction between ABTS and either potassium persulfate (Re et al., 1999) or metmyoglobin/ H2O2 (Miller et al., 1997) which was then commonly used for the screening of total antioxidant activity. In other words, the antioxidant activity of urine against ABTS+• required investigation. So, ABTS+• was generated, using the potassium persulfate method as described in 2.2.14, and mixed with 20-times diluted urine. In 2 of the 3 urine samples tested (Table 3.16; TS020207 & LH020207), there was no detectable absorbance at 730 nm, indicating 79 the complete elimination of ABTS+• by the urinary matrices. Sample MR020207, which was the least concentrated among the 3 samples (judging from the creatinine content given in Table 3.16), was able to quench almost 60% of the total pre-formed ABTS+• (estimated from the absorbance of the PBS control) within 3 minutes of mixing. Table 3.16. Quenching effect of urine on preformed ABTS+•. A solution of ABTS+• prepared as described in 2.2.14, was mixed with a fixed volume of PBS or one of the 3 urine samples (diluted 20 times with PBS) indicated below as reaction mixture content and the relative absorbance at λ=730 nm, 3 minutes after mixing, were measured. Each absorbance value is the average of duplicate readings. Reaction mixture content Relative absorbance (730 nm), 3 mins after mixing Creatinine content of urine (mM) PBS TS020207 LH020207 MR020207 1.006 < 0.001 < 0.001 0.426 - 8.18 6.96 2.85 Antioxidants in urine The source of the ABTS+• quenching effect of urine may originate from the diet, where polyphenols are the most abundant antioxidants ingested (Scalbert et al., 2005). Many polyphenolic compounds of various classes and their metabolites were detected in human urine after certain foods, especially fruits, vegetables and beverages like green tea, black tea, cocoa, grape-skin extract, wine, coffee and fruit juices were consumed (Ito et al., 2005; Mennen et al., 2006). Human subjects who participated in the urinary H2O2 studies were not restricted from taking some of the listed foods and drinks. Mammalian lignans (enterodiol and enterolactone), phenolic acids (chlorogenic, caffeic, m-coumaric, gallic, and 4-O-methylgallic acids), phloretin and flavonoids (catechin, epicatechin, quercetin, isorhamnetin, kaempferol, hesperetin, and naringenin) were excreted at a rate of 0 to 25 µmol/24hrs and could be quantified in urine by HPLC-tandem MS techniques 80 (Ito et al., 2005; Mennen et al., 2006; Tian et al., 2006). These compounds (Fig. 3.15) in urine could scavenge the ABTS+• which was formed quantitatively from urinary H2O2. Fig. 3.15. Chemical structures of polyphenols and their metabolites detected in urine (adapted from Ito et al., 2005). Another likely cause of ABTS+• suppression is ascorbic acid (AA). Arnao et al. (1996) had demonstrated that in the presence of AA, there was a delay in the generation of ABTS+• from the ABTS/H2O2/HRP system, and this lag time increased with higher concentrations of AA. Arnao et al. (1996) had also proposed the reaction as follows: AA reacts with ABTS+• to produce an ABTS and a monodehydroascorbic acid (MDHA), which reacts with one more ABTS+• to form another ABTS and a dehydroascorbic acid 81 (DHA) (Fig. 3.16b). Basically, complete reduction of one AA molecule would consume 2 ABTS+•. Note that one H2O2 molecule reacts with 2 ABTS molecules (in the presence of peroxidase) to give 2 ABTS+• (Childs et al., 1975). It had been mentioned earlier that about 0.15 mM AA could be excreted in urine which could theoretically quench 300 µM ABTS+• produced from 150 µM of H2O2 (Fig. 3.13 & 3.16). Since urinary H2O2 is usually at a lower concentration than that, it is unlikely to be detected by the ABTS assay, based on the levels of AA alone. Like AA, other antioxidants found in urine like quercetin, kaempferol and caffeic acid could react with ABTS+• within one minute of mixing (Re et al., 1999). Fig. 3.16. Mechanism of action for (a) ABTS/HRP/ H2O2 and (b) ascorbic acid (AA) with ABTS+•. MDHA, monodehydroascorbic acid; DHA, dehydroascorbic acid. A conclusion can be made at this point that ABTS is not a suitable peroxidase substrate to be used for urinary H2O2 analyses. 82 3.3.4. N-Acetyl-3,7-dihydroxyphenoxazine (amplex red) assay Background Amplex red is a non-fluorescent molecule that when oxidized by H2O2 in the presence of HRP produces resorufin, a highly-fluorescent and stable product. It has low background fluorescence but a small amount of H2O2 is sufficient to increase the fluorescence substantially (later demonstrated in Fig. 3.17). It has a high extinction coefficient (54000 M-1cm-1, 571nm; Zhou et al., 1997). An advantage of this probe over HVA and HPAA is the excitation and emission wavelengths (563/587nm) which lie in a spectral zone that has little susceptibility to interference from autofluorescence in assays of biological samples such as blood and urine (Zhou et al., 1997). The reaction stoichiometry of amplex red and H2O2 is determined to be 1:1 (Fig. 3.18; Zhou et al., 1997) and its oxidation to resorufin is an irreversible process (Reszka et al., 2005). Since the amplex red-peroxidase assay kit (A22188) from Invitrogen was not packaged in a manner and amount suitable for our purposes (refer to section 1.8), the assay was further modified to examine its suitability for urinary H2O2 measurements. Linearity Fig. 3.17 shows a typical standard calibration plot for the assay. It was found to be linear in the range of 0.2 to 10 µM, with r2 > 0.99. Any fluorescence arising from PBS + reagent solution was corrected to zero and data from the other standard concentrations were adjusted accordingly. 83 Standard Calibration Plot (Amplex Red Assay) Net Fluorescence Intensity 16000 14000 y = 1410.6x R2 = 0.9991 12000 10000 8000 6000 4000 2000 0 0 1 2 3 4 5 6 7 8 9 10 11 Concentration of H2O2 in µ M Concentration of H2O2 in µM Net Fluorescence Intensity 0 0 0.2 0.4 1 2 354 673 1466 2980 (±64) (±105) (±127) (±102) Concentration of H2O2 in µM 3 4 6 8 10 Net Fluorescence Intensity 4438 5786 8625 11285 13851 (±24) (±12) (±154) (±217) (±297) Fig. 3.17. A standard calibration plot for amplex red assay. In the above plot, 0, 0.2, 0.4, 1, 2, 3, 4, 6, 8 and 10 µM H2O2 solutions were used to calibrate for resorufin formation. Each data point is the mean ± SD of 3 separate experiments where in each experiment, duplicate measurements were made. The relative fluorescence (after correction with respect to 0 µM H2O2) using λexc/λemiss =563/587 nm as well as SD are tabulated below the plot. The relationship was found to be linear up to at least 10µM with an r2 > 0.99. Detection limit This assay was able to detect as low as 0.2 µM of H2O2, thus beating the HVA assay. Its sensitivity is attributed to the high fluorescence of the oxidized product, resorufin. This means that the assay is capable of distinguishing samples with very small 84 differences in H2O2 concentration, making it a possibly good candidate for urinary H2O2 analyses. Effect of dilution of urine samples on analyses In Table 3.17, the effect of dilution of two different urine samples obtained from the same individual but on 2 different days is shown. The H2O2 readings increased after at least 2 X dilutions. Sample RD131106, which has a higher urinary creatinine concentration, stabilized after 4 X dilutions while sample RD1411061 stabilized after approximately 2 X dilutions. These data indicate that dilutions of urine samples are necessary in further studies of this assay, in order to reduce the matrix interference while maintaining the concentrations of HRP and amplex red in the reaction mixture constant, thus increasing the probability of reaction of H2O2/HRP with amplex red rather than with other interfering compounds in urine. Table 3.17. Effect of dilution of urine sample on amplex red assay. Two urine samples were collected from the same individual but on 2 different days. Each sample was subjected to the stated number of times of dilution with PBS before analyses and the corresponding calculated concentrations of endogenous H2O2 in the undiluted sample are given in the table. Subject Urinary creatinine (mM) RD131106 RD1411061 12.2 8.13 Number of times dilution and the calculated concentration of total H2O2 (µM) Undi2X 4X 5X 10X 20X 40X 50X luted 2.44 5.83 10.3 10.4 12.5 11.6 11.6 12.3 5.98 8.63 9.98 10.3 12.0 10.3 15.3 9.16 85 Sample recovery data The neat and spiked urine samples were analyzed after the stated number of dilutions and reaction with Amplex Red/HRP (Table 3.18). All the nine samples had recovery percentages of different levels of spiked H2O2 from about 20% to 50%. No samples had any recovery above 50.5 %. Six of the urine samples were fairly dilute, containing between 2.1 to 5.8 mM creatinine and these samples were diluted 10 X with PBS. The remaining 3 samples contained 10.0, 14.6 and 15.3 mM creatinine, and these were diluted 20 X with PBS. The recovery percentages were more or less consistent throughout these nine samples but not sufficiently good for consideration as an assay for urinary analyses. All unspiked urine samples were analyzed at two dilution levels to confirm the suitability of the first chosen number of dilutions i.e. 10X dilution samples were reconfirmed with 20X dilution analyses while 20X dilutions were reconfirmed with 25X dilutions; and the calculated endogenous urinary H2O2 concentrations from both levels were comparable (data not shown). The recovery data demonstrated that even though the urine samples have been optimally diluted to reduce the concentration of interfering matrices, the amplex red assay continued to underestimate the amount of spiked H2O2. Attempting a higher number of dilutions (30 X and above) is not recommended as the diluted urinary H2O2 value may drop below the detection limit of the assay. In the next step, a comparison study of the amplex red assay with the O2 electrode assay was conducted to see if it is also underestimating endogenous urinary H2O2 concentrations. 86 Table 3.18. Amplex red assay recovery study. Urine was freshly voided from different individuals as well as the same individuals but on different days to give a total of 9 different samples. Each urine sample collected was divided into 6 portions, and each of them was spiked with different amounts of H2O2 so that after the stated number of dilutions below, they would have 0, 0.5, 1, 2, 3 and 4 µM of spiked H2O2 on top of the endogenous concentration. The recovery percentages of spiked H2O2 were calculated as described in 2.2.16. Urinary Urinary No of H2O2 creatinine dilutions (µM)# (mM) before assay 13.3 10.0 20 Concentration of spiked H2O2 (µM) and their respective recoveries (%) 0.5 1 2 3 4 38.3 42.6 29.4 28.6 34.4 Subject Sex RD1411062 M LH151106 F 2.59 2.53 10 49.6 41.5 29.0 26.8 27.8 SR151106 F 3.12 5.14 10 19.5 28.8 34.8 29.7 31.9 TS151106 M 8.31 3.10 10 50.5 21.2 46.8 33.3 43.4 RD151106 M 11.6 5.79 10 16.1 15.5 20.0 35.6 35.0 WH171106 M 23.4 15.3 20 32.6 22.4 23.8 21.1 27.4 MR171106 F 7.93 2.13 10 30.4 35.3 45.2 38.1 35.2 TS201106 M 6.89 4.96 10 47.4 46.7 34.0 37.9 40.4 RD201106 M 28.9 14.6 20 7.80 7.18 19.9 26.0 25.5 # Urinary H2O2 concentration of each sample was also determined at higher dilutions than stated above (data not shown) and the values were found to be closely similar to those given in the above table. Comparison study between amplex red and O2 electrode assays Once again, the O2 electrode assay was used as a reference method to gauge the reliability of the assay under investigation. 10 spot urine samples were freshly voided from various individuals and analyzed by the two assays within a short time interval apart. Looking at Table 3.19, all samples, with the exception of RD201106, did not have 87 closely-agreeing values between the amplex red assay and the O2 electrode assay. In all the ten samples, the amplex red assay was underestimating the endogenous urinary H2O2 concentrations. In fact, the amplex red assay readings were at least 50% smaller than the O2 electrode readings for 80% of the samples. Table 3.19. Comparison of the amplex red assay with O2 electrode assay in a few individuals. Urine samples were freshly voided from different individuals on the same as well as different days so that a total of 10 different samples were obtained. These were analyzed for H2O2 using both the amplex red assay and the O2 electrode assay. The data of both assays were derived from the mean of 2 analyses. Amplex red assay mean data for each sample was obtained using 2 different dilution levels. Urinary H2O2 in µM Amplex Red Assay O2 Electrode Assay 11.1 (± 7.5%) 34.9 (± 1.4%) Subject Sex RD1411061 M RD1411062 M 13.4 (± 0.3%) 31.5 (± 3.1%) LH151106 F 2.67 (± 3.0%) 19.3 (± 2.2%) SR151106 F 2.97 (± 5.1%) 15.9 (± 2.7%) TS151106 M 8.20 (± 1.3%) 22.7 (± 1.9%) RD151106 M 11.6 (± 0.1%) 30.0 (± 2.9%) WH171106 M 23.0 (± 1.9%) 35.1 (± 2.4%) MR171106 M 7.79 (± 1.7%) 21.0 (± 2.0%) TS201106 M 6.48 (± 6.3%) 12.8 (± 2.0%) RD201106 M 29.4 (± 1.7%) 32.7 (± 3.9%) Failure of the amplex red assay The oxidation product of amplex red, resorufin, is also a substrate of HRP and can be further oxidized to a non-fluorescent compound, resazurin (Fig. 3.18). However, this further oxidation will not occur significantly unless the H2O2 concentration is higher than the amplex red concentration in the reaction mixture (Zhou et al., 1997). Such a situation 88 is unlikely to happen in this assay and is not the cause of the earlier observed underestimations; even when the highest H2O2 standard concentration (10 µM) is employed, its initial concentration in the reaction mixture is only 2.5µM as compared to amplex red at 120µM. Fig. 3.18. HRP-catalyzed amplex red oxidation by H2O2 (adapted from Towne et al., 2004) Towne et al. (2004) found that in the absence of HRP and H2O2, discernible changes in the fluorescence intensity of resorufin were observed by simply dissolving resorufin in aqueous solutions in the pH range 6.2 to 7.7, near its pKa value of 6.5. This observation could be due to de-N-acetylation and nucleophilic addition, leading to polymerization of resorufin (Fig. 3.18, Reaction 2). Loss of resorufin fluorescence by this mechanism could not be the source of underestimation in my studies, although the reaction mixtures were buffered with PBS to be in that region of pH. This was because 89 Towne et al. (2004) found that the non-enzymatic decay of fluorescence intensity in pH6.5, 0.05M phosphate buffer was lowest for 10 µM resorufin, remaining fairly stable for at least 60 minutes and it only became pronounced at higher resorufin concentrations of 20-160 µM. Less than 10 µM resorufin would be formed in the assay. These data suggest that the oligomerization or polymerization of resorufin occur with higher resorufin concentration. Incubation time in this assay was kept as short as possible (2 min) to minimize any contribution from reactions other than the intended Reaction 1 (Fig. 3.18, Towne et al., 2004). So, why is the amplex red assay still found to be not suitable for the determinations of urinary H2O2 concentrations? Or what actually caused the poor recovery of spiked H2O2 from urine in this assay? The answer lies in the possible interferences coming from other peroxidase substrates present along with amplex red in the complex biological sample of human urine. They may compete with amplex red for the enzyme while consuming H2O2. Accordingly, less amplex red will be converted to resorufin, and this will underestimate the actual level of peroxide (Reszka et al., 2005). Anticancer anthracenediones (mitoxantrone and ametantrone) and common analgesics (e.g. acetaminophen) can inhibit the formation of resorufin from amplex red at low H2O2 concentration (Fig. 3.19; Reszka et al., 2005), but they are unlikely to be present in the urine of healthy subjects. p-Hydroquinone (Fig. 3.19) and nitrites are good peroxidase substrates too (Reszka et al., 2005); the former can be found in the urine of benzeneexposed workers (Kim et al., 2006) while the latter can be found in normal urine (Pannala et al., 2003) and especially higher in urine of subjects with urinary tract infections (Lundberg et al., 1997). Ascorbate is only one of many possible interfering 90 compounds but a more likely candidate to be found in urine of subjects which can have an inhibitory effect on HRP-based analyses (Martinello et al., 2006). Fig. 3.19. Structures of some peroxidase substrates (adapted from Reszka et al., 2005). Thus, the amplex red was not suitable as a probe for urinary H2O2 due to its inability to compete effectively with other peroxidase substrates present in human urine for HRP/ H2O2. 91 3.3.5. 2’,7’-Dichlorodihydrofluorescein (DCFH) assay Background In the presence of HRP/ H2O2, non-fluorescent 2’,7’-dichlorodihydrofluorescein (DCFH) is oxidized to fluorescent 2’,7’-dichlorofluorescein (DCF) (Fig. 4.1). Like amplex red, it also has low background fluorescence but a small amount of H2O2 is sufficient to increase the fluorescence substantially (Fig. 3.20). In fact, its response is at least 3 times greater than amplex red, which makes it attractive to study in this section. Linearity Fig. 3.20 shows a typical standard calibration plot for the assay. It was found to be linear in the range of 0.2 to not more than 10 µM, with r2 > 0.99. Any fluorescence arising from PBS + DCFH reagent solution was corrected to zero and data from the other standard concentrations were adjusted accordingly. Detection limit Like the amplex red assay, this one could also detect as low as 0.2 µM of H2O2. Its sensitivity is attributed to the high fluorescence of the oxidized product, DCF. The assay thus has the potential of being able to distinguish samples with very small differences in H2O2 concentration, making this a possibly good candidate for urinary H2O2 analyses. 92 Net Fluorescence Intensity Standard Calibration Plot (DCFH Assay) 50000 45000 y = 4430.7x 40000 35000 R 2 = 0.9955 30000 25000 20000 15000 10000 5000 0 0 2 4 6 8 10 12 Concentration of H2O2 in µ M Concentration of H2O2 µM Net Fluorescence Intensity Concentration of H2O2 µM Net Fluorescence Intensity 0 0 0.2 1008 (±90) 0.5 2458 (±151) 1 4954 (±432) 2 9933 (±587) 3 14575 (±989) 4 5 6 7 8 10 18722 23194 27670 30794 35266 42369 (±1523) (±1662) (±1827) (±2158) (±2897) (±2983) Fig. 3.20. A standard calibration plot for DCFH assay. In the above plot, 0, 0.2, 0.5, 1, 2, 3, 4, 5, 6, 7, 8 and 10 µM H2O2 solutions were used to calibrate for DCF formation. Each data point is the mean ± SD of 3 separate experiments where in each experiment, duplicate measurements were made. The relative fluorescence (after correction to zero with respect to 0 µM H2O2) using λexc/λemiss =498/522 nm as well as SD are tabulated below the plot. The relationship was found to be linear up to a maximum of 10µM with an r2 > 0.99. Effect of dilution of urine samples on analyses In Table 3.20, the effect of dilution of 5 different urine samples collected on different days from 3 individuals is shown. In all cases, the urinary H2O2 measurements were lowest when there was no dilution of the urine samples, as much as 52% less than the value obtained with a 2 X dilution. It was noted that the H2O2 concentration values 93 were stable and reproducible at 4 (or 5), 10 and 20 X dilutions for each sample. In fact, sample TS190606, which had the lowest urinary creatinine concentration among the five samples (at 3.98 mM), stabilized after just 2X dilutions while the most concentrated sample RD130606 (with 14.9 mM creatinine) stabilized after 5X dilutions. Thus, as in the Amplex Red assay, a higher urine concentration would require a higher number of dilutions prior to reaction, to remove as much of the sample interferences as possible. Table 3.20. Effect of dilution of urine sample on DCFH assay. Urine was freshly voided from different individuals as well as same individuals but on different days to give a total of 5 different samples. Each sample was subjected to the stated number of times of dilution with PBS before analyses and the corresponding calculated concentrations of endogenous H2O2 in the undiluted sample are given in the table. The graph below helps to illustrate the change of calculated [H2O2] with increasing number of dilutions. Urinary Number of times dilution and the calculated concentration of creatinine total H2O2 (µM) (mM) Undiluted 2X 4X 5X 10X 20X RD130606 14.9 8.92 17.9 19.9 22.9 22.7 20.7 RD190606 12.4 7.04 14.9 19.3 nd 18.3 17.51 TS190606 3.98 11.0 15.4 16.3 nd 16.1 19.0 RD210606 8.87 10.4 16.8 19.2 nd 20.3 21.7 WH210606 4.41 9.44 18.1 23.2 nd 25.0 26.3 nd : not determined in experiment Calculated Conc of H 2O2 in µ M Subject 30 25 20 15 10 5 0 1 2 4 10 20 No of times dilution of urine samples 94 Recovery study data The neat and spiked urine samples were analyzed and the calculated recovery percentages are tabulated in Table 3.21. The 14 different urine samples collected varied in their concentrations, as indicated by their creatinine content, ranging from the most dilute at 1.46 mM to the most concentrated at 24.9 mM. These samples were diluted between 4 to 20 X after spiking and before reacting with DCFH/HRP. All the samples generally gave good recovery percentages with different levels of spiked H2O2. The results obtained were by far the best among all HRP-based assays tested. They also surpassed all chemical-based methods, except the O2 electrode and to a certain degree, the FOX-2 assay. In the last 9 samples listed in the table, two sets of recovery percentages were given; one set is the result of 30 minutes of incubation with DCFH/HRP and the other (in parenthesis) with only 10 minutes. The difference between the two sets is relatively small, with more than 90% of the data differing by only 0.3 – 6.0 % in recovery percentage between the two incubation times. All samples (except for RD140806 at 2 levels; 0.5 & 1 µM) had more than 60% of the spiked H2O2 recovered in calculation. In fact, 9 out of 14 samples had recoveries of more than 80%. It is highly possible that these recovery percentages could be much further improved for some samples in the lower half of the table by increasing the number of dilutions of the sample (to 10, 15 or 20 X). Since the required number of dilutions was approximated by looking at the intensity of colouration of the urine, samples in further DCFH assay experiments were analyzed using at least 2 different dilution levels to check for reproducibility of data and to show that the number of dilutions chosen was acceptable. 95 Table 3.21. DCFH assay recovery study. Urine was freshly voided from different individuals as well as the same individuals but on different days to give a total of 14 different samples. Each urine sample collected was divided into 6 portions, and each of them was spiked with different amounts of H2O2 so that after the stated number of dilutions below, they would have 0, 0.5, 1, 2, 3 and 4 µM of spiked H2O2 on top of the endogenous concentration. The recovery percentages of H2O2 spiked were calculated as described in 2.2.16. (*) Values indicated in parenthesis were derived from 10 min of incubation with DCFH reagent solution while others were from 30 min of incubation. Subject Sex *Urinary Urinary No of H2O2 creatinine dilutions (µM) (mM) before assay 39.1 20.6 5 RD140606 M TS150606 M 13.0 2.01 RD210606 M 26.1 WH210606 M RD150606 *Concentration of spiked H2O2 (µM) and their respective recoveries (%) 0.5 1 2 3 4 102.6 103.7 112.9 115.4 113.8 10 96.3 96.0 88.3 92.4 89.2 8.87 15 114.6 97.4 92.0 95.0 90.3 36.2 4.41 15 79.6 91.8 79.0 89.9 87.6 M 41.4 24.9 20 122.1 127.6 116.9 115.9 113.8 LZ080806 F 4.12 (4.12) 1.46 4 137.1 108.3 101.1 95.1 88.1 (117.7) (103.3) (97.7) (91.8) (87.8) SR110806 F 6.72 (6.60) 4.58 4 84.3 (83.4) 90.3 (88.9) 89.1 94.4 92.8 (85.6) (92.3) (90.4) RD070806 M 17.2 (16.3) 4.83 4 74.8 (67.1) 77.1 (66.1) 71.7 74.6 66.3 (65.9) (68.6) (61.5) LH110806 F 2.42 (2.46) 2.68 4 66.2 (61.0) 63.8 (60.5) 72.1 71.5 69.9 (68.3) (70.7) (68.2) LZ090806 F 20.0 (18.7) 14.7 8 63.5 (61.9) 58.9 (58.2) 54.5 65.1 71.9 (52.8) (60.4) (67.9) RD140806 M 48.9 (46.8) 12.7 8 48.6 (50.4) 49.5 (45.9) 70.1 63.9 67.8 (66.9) (63.0) (64.4) GJ080806 M 23.5 (22.6) 9.03 10 106.1 (100.1) 92.2 (94.2) 87.3 93.4 84.9 (82.1) (87.0) (80.2) WH140806 M 31.3 (29.7) 17.3 10 91.3 (89.5) 66.3 (64.8) 71.5 63.7 65.4 (72.0) (62.7) (64.1) GJ110806 M 19.6 (18.2) 11.3 10 72.1 (77.8) 111.6 101.1 96.9 90.2 (110.7) (96.0) (92.9) (88.6) 96 10 µM 45000 40000 8 µM 35000 6 µM 30000 5 µM 25000 4 µM 20000 3 µM 15000 2 µM 10000 1 µM 5000 0 µM 0 0 Vmax Points = 29 100 200 300 400 500 600 700 Time (secs) Fig. 3.21. Fluorescence intensity progress of the DCFH/HRP reaction with 0 to 10 µM H2O2 standards. Reaction for the lowest concentration was instantaneous while reaction for the highest standard concentration was completed by 10 mins. Maximum time required for reaction completion When working standards of between 0 to 10 µM H2O2 were reacted with the DCFH reagent solution, the reaction progress chart in Fig.3.21 was obtained. Reactions with 1 and 2 µM H2O2 were instantaneous. The highest standard concentration (10 µM) completed its reaction by 10 minutes. Thus, it was not necessary to incubate the reaction up to 30 minutes. 97 Data differences between 10-min and 30-min incubation The differences in the recovery percentages of spiked H2O2 between the two incubation times in Table 3.21 had been earlier mentioned to be relatively small. Similarly, the endogenous urinary H2O2 did not vary much between the two incubation times in Table 3.21. Sample LZ080806 had negligible difference (about 4.12 µM for both incubation times) while sample GJ110806 had its 10-min incubation value within 7.1% of its 30-min incubation value (18.2 and 19.6 µM respectively). Other samples had their differences lying between 1.6 and 5.2% of the 30-min incubation value. For most of the samples, the 30-min incubation values were slightly higher than the 10-min incubation values. The more concentrated the urine, the greater the difference between the two incubation times but this difference gets even smaller with more dilutions of the urine sample (data not shown). Comparison study between DCFH and O2 electrode assays Once again, the O2 electrode assay was used as a reference method to gauge the reliability of the assay under investigation. 19 spot urine samples were freshly voided from various individuals and analyzed by the two assays in parallel. Looking at Table 3.22, the DCFH assay and the O2 electrode assay gave comparable urinary H2O2 readings for about 85% of the samples. Generally, the two assays showed very good agreement with each other, surpassing the performance of other comparison studies with the O2 electrode assay, involving the FOX-2 and amplex red assays. 98 Table 3.22. Comparison of the DCFH assay with O2 electrode assay in a few individuals. Urine samples were freshly voided from different individuals as well as the same individuals but on different days so that a total of 19 different samples was obtained. These were analyzed for H2O2 using both DCFH assay and O2 electrode assay. DCFH assay data were derived either from the mean of 3 dilution levels’ determinations (± SD) or from the average of duplicate fluorescence measurements at one dilution level, while O2 electrode assay data were the mean of 2 analyses (± % difference). DCFH assay data with ± SD were obtained through 30-min incubation while the rest were through 10-min incubation. Urinary H2O2 in µM DCFH Assay O2 Electrode Assay 18.4 ± 0.9 8.98 (± 8.4%) Subject Sex RD190606 M TS 190606 M 17.2 ± 1.6 19.5 (± 7.7%) RD210606 M 20.4 ± 1.2 13.7 (± 2.4%) WH210606 M 24.8 ± 1.5 25.4 (± 0.0%) LH260606 F 19.3 ± 0.6 21.9 (± 6.7%) SR260606 F 12.9 ± 1.0 7.78 (± 12.5%) SW260606 F 8.74 ± 0.9 6.32 (± 23.0%) RD260606 M 25.5 ± 0.3 23.3 (± 4.2%) LZ260606 F 11.5 ± 0.7 13.1 (± 3.7%) KL260606 F 15.0 ± 3.4 9.82 (± 1.0%) RD070806 M 16.3 18.8 (± 3.3%) LZ080806 F 4.12 7.21 (± 4.3%) LZ090806 F 18.7 6.59 (± 23.8%) GJ080806 M 22.6 18.8 (± 0.0%) LH110806 F 2.46 3.74 (± 19.9%) SR110806 F 6.60 8.60 (± 4.4%) GJ110806 M 18.2 26.2 (± 14.3) WH140806 M 29.7 21.9 (± 0.0 %) RD140806 M 46.8 28.2 (± 0.0 %) 99 Table 3.23. Coefficient of variation of various urine samples analyzed by DCFH assay. Sample RD191206 was analysed at 10 and 20 X dilutions, and 4 repeats were made at each dilution level, giving n = 8. Other samples in the table were analysed at (4 or 5), 10 and 20 X dilutions, giving n = 3. The corresponding SD and CV were calculated. Urine Sample RD191206 RD190606 TS 190606 RD210606 WH210606 LH260606 SR260606 SW260606 RD260606 LZ260606 Mean conc. of H2O2 in µM 54.6 18.4 17.2 20.4 24.8 19.3 12.9 8.74 25.5 11.5 n 8 3 3 3 3 3 3 3 3 3 SD 3.4 0.9 1.6 1.2 1.5 0.6 1.0 0.9 0.3 0.7 CV% 6.1 4.9 9.3 5.9 6.0 3.1 7.8 10.3 1.2 6.1 Coefficient of variation There was good agreement between repeats for every sample analyzed by the DCFH assay, with the coefficient of variation (CV) around 10 % or less (Table 3.23). Study of the effect of catalase and SOD on the DCFH reaction fluorescence Two urine samples were collected from two subjects, diluted 2X and treated with either PBS only, catalase or superoxide dismutase (SOD). The reaction progress as monitored by their fluorescence at 522nm was shown in Figure 3.22. The urine reaction mixture with no additional treatment had the highest fluorescence throughout the reaction time monitored (trace A). When 20 or 40 U of SOD was added, the fluorescence intensity fell by a maximum of 18% of the no-treatment mixture (at 30 min). When any amount of catalase or a mixture of catalase and SOD was added, the fluorescence intensity dropped by as much as 74% (at 10 min). 100 The only slight drop in fluorescence when SOD was added to urine showed that the DCFH/HRP system reacts mainly with urinary H2O2, and not O2•-. The small reduction could perhaps be attributed to the removal of a small amount of H2O2 that was generated through autooxidation of compounds like polyphenolics in urine, involving O2•. The substantial abolishment of fluorescence signal upon the addition of catalase further reinforced the specificity of the DCFH assay to urinary H2O2. DCFH: a reliable HRP substrate for assay of urinary H2O2 Further discussion and explanation of the results of the DCFH assay studies is presented in Chapter 4. At this point of time, it would suffice to say that the DCFH assay is suitable for the purposes of urinary H2O2 investigations based on its high sensitivity, specificity, very low detection limit, linearity of response, good coefficient of variation (reproducibility), good recovery of spiked H2O2 and its close agreement with the O2 electrode assay. 101 20000 19000 18000 17000 16000 S1 15000 A 14000 13000 B 12000 C 11000 10000 9000 8000 7000 D, F, E 6000 5000 4000 3000 2000 1000 0 0 100 200 300 400 500 600 700 800 900 1000 1100 1200 1300 1400 1500 1600 1700 1800 1900 2000 2100 1900 2000 2100 Time (secs) Vmax Points = 8 A 39000 B S2 34000 C 29000 24000 19000 D 14000 E F 9000 4000 0 100 200 300 400 500 600 700 800 900 1000 1100 1200 1300 1400 1500 1600 1700 1800 Time (secs) Vmax Points = 8 Fig. 3.22. Fluorescence intensity progress of the DCFH/HRP reaction in 2 urine samples (S1 & S2). Each freshly-voided urine sample was diluted 2X. 1 ml portions of each diluted urine sample were given one of the following treatments: Trace A, PBS; trace B, 40U SOD; trace C, 20U SOD; trace D, 40U catalase; trace E, 20U catalase; trace F, 20U catalase + 20U SOD. The horizontal axis indicates the time of reaction while the vertical axis gives the relative fluorescence intensity. 102 3.3.6. Dihydrorhodamine 123 (DHR) assay Background Dihydrorhodamine 123 (DHR) has a chemical structure similar to DCFH, except that while DCFH has a fluorescein backbone (Fig. 4.1 & 4.4), the 3’ and 6’ positions in DHR are substituted with amino groups while its 2’ and 7’ positions have no substituent groups (Fig. 3.23). Thus, it will be interesting to see if DHR may share the same success as DCFH in determining urinary H2O2 in the presence of HRP. DHR is a non-fluorescent molecule that is oxidized by H2O2/HRP to rhodamine 123 (Henderson et al., 1993), a cationic probe which has a higher fluorescence than DCF. The molar extinction coefficients at 500 nm for DCF and rhodamine 123 are 59,500 and 78,800 M-1cm-1 respectively (Crow, 1997). Fig. 3.23. Oxidation of dihydrorhodamine 123 (DHR) to rhodamine 123 (adapted from Crow, 1997). The first notable difference of DHR was its poor solubility in aqueous systems, unlike DCFH. Methanol formed 30% by volume of the reagent solution to counter this 103 problem. This concentration of added methanol did not affect the function of HRP (Ryu et al., 1992) and the outcome of the assay. The background fluorescence coming from the blank reaction mixture is a lot higher than that for amplex red or DCFH but the response to H2O2 was at least 3 times higher than that of DCFH for the same concentration of H2O2 (Fig. 3.24). Standard Calibration Plot (DHR Assay) Net Fluorescence Intensity 120000 y = 17118x R2 = 0.9995 100000 80000 60000 40000 20000 0 0 1 2 3 4 5 6 7 Concentration of H2O2 in µ M Concentration of H2O2 µM Net Fluorescence Intensity Concentration of H2O2 µM Net Fluorescence Intensity 0 0 0.5 1 2 8416 17000 34565 (±587) (±861) (±1500) 3 4 5 6 53224 68130 86017 101567 (±2340) (±2950) (±7090) (±11100) Fig. 3.24. A standard calibration plot for DHR assay. In the above plot, 0, 0.5, 1, 2, 3, 4, 5 and 6 µM H2O2 solutions were used to calibrate for rhodamine 123 formation. Each data point is the mean ± SD of 3 separate experiments where in each experiment, duplicate measurements were made. The relative fluorescence (after correction to zero with respect to 0 µM H2O2) using λexc/λemiss =505/529 nm as well as SD are tabulated below the plot. The relationship was found to be linear up to a maximum of 6µM with an r2 > 0.99. 104 Linearity Fig. 3.24 shows a typical standard calibration plot for the assay. It was found to be linear in the range of 0.5 to not more than 6 µM, with r2 > 0.99. Beyond the last point, the fluorescence was too high to obey a linear relationship with the concentration of H2O2. The blank reaction mixture was corrected to zero fluorescence and data from the other standard concentrations were adjusted accordingly. Detection limit Although rhodamine 123 has a higher fluorescence than DCF, the DHR assay was not as sensitive to low amounts of H2O2 as the DCFH assay. The detection limit was slightly higher at 0.5 µM of H2O2. Samples data: effects of dilution The fluorescence emitted from reaction mixtures with urine was slightly lower than that obtained with the blank reaction mixture. Table 3.24 shows that the calculated urinary H2O2 values were unrealistically low and did not stabilize with more dilutions; the values kept increasing but very minimally. The DHR assay was also compared with the DCFH assay for two of the urine samples, RD121006 and RD131006. With the DCFH assay, the former sample gave reproducible readings at 10, 15 and 20X dilutions while the latter gave fairly stable readings at 5, 10 and 20X dilutions. 105 Table 3.24. Effect of dilution of urine sample on DHR assay and comparison with DCFH assay. Urine was freshly voided from the same individual but on different days to give a total of 3 different samples. Each sample was subjected to the stated number of times of dilution with PBS before analyses and the corresponding calculated concentrations of endogenous H2O2 in the undiluted sample were given in the table. 2 of the 3 samples were similarly diluted and analyzed by the DCFH assay for purposes of comparison. Subject Assay RD111006 DHR DHR DCFH RD121006 Assay Subject RD131006 DHR DCFH Number of times dilution and the calculated concentration of total H2O2 (µM) Undiluted 2X 4X 8X 10X 15X 20X 0.12 0.26 0.37 1.81 1.75 3.35 3.84 negative 0.04 0.06 0.33 negative values 16.8 32.8 59.5 74.4 79.2 79.8 81.3 Number of times dilution and the calculated conc. of total H2O2 (µM) Undiluted 5X 10X 20X 0.02 0.50 0.59 1.39 13.3 23.2 25.4 26.2 Samples data: recovery study Spiking different amounts of H2O2 into separate portions of just one sample was enough to show that the DHR assay was not suited for urinary H2O2 analyses. Recovery percentages range miserably between 1.9 to 5.4% (Table 3.25). The same neat and spiked portions were analyzed simultaneously by the DCFH assay and the recovery percentages were in the good range of 77.0 to 83.2% (Table 3.25). 106 Table 3.25. DHR assay recovery study and comparison with DCFH assay. One spot urine sample was collected and divided into 6 portions, and then spiked with different amounts of H2O2 so that after the stated number of dilutions below, they would have 0, 0.5, 1, 2, 3 and 4 µM of spiked H2O2 on top of the endogenous concentration. The recovery percentages of spiked H2O2 were calculated as described in 2.2.16. Subject Urinary Creatinine (mM) RD131006 9.48 Assay DHR DCFH Concentration of spiked H2O2 Urinary No of (µM) and their respective H2O2 dilutions recoveries (%) (µM) before assay 0.5 1 2 3 4 1.09 10 5.4 4.9 2.8 2.5 1.9 35.1 10 78.0 77.0 82.8 83.2 82.1 Differences in reaction of DCFH and DHR The difference in substituent group turned out to make DCFH and DHR worlds apart in their performances in urinary H2O2 determination. The DHR assay failed to detect realistic levels of H2O2, even after sample dilutions of up to 20X (Table 3.24). The assay also failed to pick up most of the H2O2 spiked into the urine samples (Table 3.25). On the other hand, the DCFH assay was able to deliver consistent values above a certain number of dilutions and the spiked H2O2 was recovered at 77.0% and above, when the same urine samples were studied in parallel, using the same number of dilutions and spiked amount of H2O2 (Tables 3.24 & 3.25). So, the problem lies with the DHR assay, and not the samples. 107 Fig. 3.25. Mechanism for the oxidation of dihydrorhodamine 123 (DHR; RH = rhodamine 123; adapted from Kooy et al., 1994) One possible reason for the failure of the DHR assay is that DHR is much less successful than DCFH in competing with other peroxidase substrates in urine for the active site of the HRP enzyme intermediate. The different functional groups on DHR and DCFH could greatly influence their relative ease of oxidation by HRP-compound I, where the removal of one electron from the substrate is likely to be the rate-determining step (Job et al., 1976; Fig. 3.25). A second possibility is the presence of urinary compounds that inhibit the reaction pathway taken by DHR to form rhodamine 123. For instance, urate might effectively reduce the DHR radicals formed (DHR•-) back to DHR (Kooy et al., 1994; Fig. 3.25). DHR is used in the study of intracellular ROS because its oxidation brings about the tautomerization of one of its two equivalent amino groups to a charged imino (Fig. 3.23), which is stably localised within the cell (Royall et al., 1993). However, it cannot be said with certainty that the cation would remain similarly stable in the urinary environment without being attacked by other species. Based on the urine dilution data and the poor recovery of spiked H2O2, the DHR assay did not look suitable for measuring urinary H2O2. 108 3.4. BASAL URINARY HYDROGEN PEROXIDE MEASUREMENTS As mentioned in the earlier sections, the DCFH assay and the O2 electrode assay were by far, the two most specific and robust assays studied for urinary H2O2 measurements. Thus, these two assays were employed to measure basal H2O2 levels in a range of healthy human subjects at different times of the day and on different days within a period of 6 months (Table 3.26). Due to variations in the water content of the collected urine samples which would influence the H2O2 concentration reading, all readings were standardized against the concentration of creatinine in the respective samples. Even after normalization, the subjects still showed a wide range of starting urinary H2O2 concentrations (at 1100hrs) within each assay (O2 electrode assay: 1.12 to 5.44 µM H2O2 per mM creatinine; DCFH assay: 0.96 to 3.33 µM H2O2 per mM creatinine). This finding was consistent with a previous study involving healthy individuals (Halliwell et al., 2004a). The variability could conceivably be related to differences in the diets of the subjects as well as to differences in endogenous rates of H2O2 generation and H2O2 catabolism (Long et al., 1999b & 2000). Physical activity and electrolyte balance within the body were also reported to affect urinary H2O2 excretion (Kuge et al., 1999). Simply put, the subjects’ lifestyle and unique individual genetic make-up could be the major influencing factors for the wide variability within each assay. Thus, the basal measurements of H2O2 for each subject were expressed as fold changes with respect to the first sample collection at 1100hrs taken as 1.00. Between the two assays, some small differences in urinary H2O2 measurements for the same samples were noted (Table 3.26). These small differences are likely due to 109 the lack of sensitivity of the O2 electrode assay, and the varying physical and chemical properties of the different urine samples (such as solute composition and pH) that could affect the accuracy of measurement by both assays, to some extent. Some of these factors will be discussed in Chapter 4. Nevertheless, data from both the O2 electrode and DCFH assays are generally quite similar to each other (Table 3.26). Based on the range of fold changes observed for each subject, the DCFH assay gave more stable base level H2O2 readings than the O2 electrode assay throughout the day for all subjects with the exception of LZ whose samples had similar magnitude of fluctuation in both assays. For the DCFH assay, the H2O2 readings did not exceed or drop by more than 50% of the initial value at 1100hrs. On the other hand, fold changes can change by more than 50% in the O2 electrode assay and was observed in 3 subjects (RD, TS and WH); for instance, contributions from subject WH at 1500 and 1700hrs were 2.62 and 2.26 folds higher than the initial value at 1100hrs, respectively. Based on the SDs, there was also generally less variation for each subject between the 3 days in the DCFH assay than in the O2 electrode assay. The newly-developed DCFH assay for urinary H2O2 measurements shows that human subjects do excrete a basal level of H2O2 that remains fairly stable throughout the day and also between the days, although the actual basal concentrations vary between individuals. More importantly, this study shows the potential suitability of urinary H2O2 as a biomarker of oxidative stress in the human body since the urinary H2O2 measured by the DCFH assay did not vary widely in the same subjects under the same conditions at different times. 110 Table 3.26. Variations in H2O2 level throughout the day as measured by two assays (DCFH and O2 electrode). Urinary H2O2 concentrations were standardized against creatinine concentrations. The data are expressed as fold changes ± SD based on the first sample collected at 1100 hrs, due to a wide variation in urinary H2O2 levels between subjects. Experiments on each subject were repeated over 3 separate days within 6 months. No coffee drinkers participated in the study. DCFH Assay Subject Sex RD SR LZ JG LH TS SW WH M F F M F M F M Initial [H2O2] (µM per mM of creatinine) 1100 hrs 1.37 ± 0.70 0.96 ± 0.16 1.81 ± 0.24 2.94 ± 1.48 1.40 ± 0.49 2.22 ± 1.13 3.33 ± 1.08 3.04 ± 1.19 No. of fold changes in [H2O2] 1100hrs 1.00 ± 0.51 1.00 ± 0.17 1.00 ± 0.13 1.00 ± 0.50 1.00 ± 0.35 1.00 ± 0.51 1.00 ± 0.32 1.00 ± 0.39 1300hrs 0.86 ± 0.26 0.92 ± 0.28 0.97 ± 0.08 0.78 ± 0.39 1.07 ± 0.35 0.98 ± 0.73 0.92 ± 0.13 1.08 ± 0.44 1500hrs 1.02 ± 0.38 1.03 ± 0.57 0.88 ± 0.20 0.96 ± 0.60 1.16 ± 0.33 0.81 ± 0.31 0.95 ± 0.40 0.97 ± 0.43 1700hrs 0.90 ± 0.33 1.21 ± 0.61 0.74 ± 0.23 0.80 ± 0.39 1.50 ± 0.37 0.74 ± 0.29 1.02 ± 0.55 1.25 ± 0.47 O2 Electrode Assay Subject Sex RD SR LZ JG LH TS SW WH M F F M F M F M Initial [H2O2] (µM per mM of creatinine) 1100 hrs 1.12 ± 0.29 1.51 ± 0.39 3.58 ± 1.62 1.90 ± 0.26 2.47 ± 1.71 4.07 ± 2.40 5.44 ± 2.21 1.80 ± 0.19 No. of fold changes in [H2O2] 1100hrs 1.00 ± 0.26 1.00 ± 0.26 1.00 ± 0.45 1.00 ± 0.14 1.00 ± 0.69 1.00 ± 0.59 1.00 ± 0.41 1.00 ± 0.11 1300hrs 1.07 ± 0.33 0.85 ± 0.34 1.01 ± 0.33 1.13 ± 0.88 1.31 ± 0.86 0.75 ± 0.69 1.04 ± 0.40 1.98 ± 0.94 1500hrs 1.13 ± 0.62 1.03 ± 0.71 0.86 ± 0.34 0.63 ± 0.11 0.66 ± 0.40 0.44 ± 0.21 0.72 ± 0.30 2.62 ± 1.41 1700hrs 1.57 ± 1.11 1.23 ± 1.31 1.12 ± 0.14 1.14 ± 1.16 0.78 ± 0.22 0.49 ± 0.26 0.64 ± 0.69 2.26 ± 0.62 111 3.5. EFFECT OF COFFEE ON BASAL URINARY HYDROGEN PEROXIDE Drinking coffee has been claimed to raise the levels of H2O2 in urine within less than 2 hours (Hiramoto et al., 2002; Long et al., 2000). In this study, the DCFH assay was used to see if this result could be confirmed. The O2 electrode assay was also employed for comparison. On a separate day, subject TS drank coffee immediately after the first collection of urine at 1100hrs. Due to the diuretic effect of coffee, subject was capable of contributing 3 further urine samples at 1130, 1200 and 1230hrs, in addition to the regular collections at 1300, 1500 and 1700hrs. Table 3.27 illustrates the effect of coffee taking on both the O2 electrode and DCFH assay readings. Within each assay, the first collection’s H2O2 concentrations for the ‘basal’ (without coffee) and ‘coffee’ days were very similar to each other. After coffee-taking at 1100-1105hrs, urinary H2O2 excretion started to increase at the second collection (1130hrs) and the values peaked at either 1200hrs (for O2 electrode assay) or 1230hrs (for DCFH assay). The H2O2 values started to decrease after the peak and came closer to the basal readings, at least by 1500 and 1700hrs. For DCFH assay, the ‘coffee’ day readings became near basal starting at 1300hrs. Even though there were no collections made between 1100 to 1300hrs on ‘basal’ days, a line joining the 1100hrs value with the 1300hrs value can be drawn to better illustrate the basal H2O2 concentrations excreted within the day, in contrast to the spike produced with coffee treatment. Hence, this experiment shows that the DCFH assay is capable of reproducing the results of other authors and verifying that coffee consumption brings about an increase in the H2O2 concentration, most likely due to additional production of H2O2 from 112 the autooxidation of hydroxyhydroquinone excreted in urine (Hiramoto et al., 2002; Halliwell et. al., 2004a; Long et al., 2000). Table 3.27. Effect of coffee consumption on urinary H2O2 concentration. The H2O2 values were standardized against creatinine. Due to the variation in initial [H2O2] as seen below, the rest of the data are presented as fold-changes. Data are mean ± SD (n = 3). For the ‘coffee’ data, subject TS consumed coffee immediately after the first urinary collection at 1100hrs on 3 separate days. For the ‘basal’ data, 3 separate experiments were repeated on subject TS without any coffee consumption. DCFH Assay Basal Coffee O2 Electrode Basal Coffee Subject TS Time Initial [H2O2] (µM per mM of creatinine) 1100 hrs 2.22 ± 1.13 2.24 ± 0.99 4.07 ± 2.40 3.87 ± 1.01 No. of fold changes in [H2O2] 1100 hrs 1130hrs 1200hrs 1230hrs 1300hrs 1500hrs 1700hrs 1.00 ± 0.51 0.98 ± 0.73 0.81 ± 0.31 0.74 ± 0.29 1.00 ± 0.44 1.34 ± 0.47 1.43 ± 0.37 1.78 ± 0.32 1.39 ± 0.42 1.01 ± 0.21 0.84 ± 0.27 1.00 ± 0.59 0.75 ± 0.69 0.44 ± 0.21 0.49 ± 0.26 1.00 ± 0.26 1.33 ± 0.58 1.90 ± 0.87 1.42 ± 0.05 0.87 ± 0.17 0.54 ± 0.01 0.43 ± 0.09 Subject TS on Coffee 2.00 µ M H2O2 per mM creatinine 1.80 1.60 1.40 1.20 DCFH Assay (Basal) DCFH Assay (Coffee) O2 Electrode (Basal) O2 Electrode (Coffee) 1.00 0.80 0.60 0.40 0.20 0.00 1100 1130 1200 1230 1300 1500 1700 Time of collection of urine sample 113 CHAPTER 4 FURTHER DISCUSSION Oxygen electrode assay The O2 electrode assay is a catalase-based electrochemical method that indirectly measures hydrogen peroxide. Urinary hydrogen peroxide in the chamber medium is first catalytically decomposed by catalase. Some of the oxygen evolved will diffuse across the membrane, through the KCl electrolyte, to the electrodes where it participates in the following electrochemistry: Silver anode: Ag → Ag+ + e-, followed by Ag+ + Cl- → AgCl Platinum cathode: O2 + 2H2O + 2e- → H2O2 + 2OH-, followed by H2O2 +2e- → 2OHA higher urinary hydrogen peroxide concentration is translated to a higher resultant O2 concentration in the chamber after catalytic decomposition. More O2 will then diffuse to the electrodes. In agreement with Faraday’s First Law of Electrolysis, a higher O2 consumption at the cathode results in a proportionally higher magnitude of current flow through the electrodes, so that a greater deflection is recorded on the chart (Fig. 3.1). Such a method as described is not likely to be affected by much interference. Even though H2O2 extraneous to the one being measured is produced through the cathode half-reaction and catalase may appear to possibly further complicate the matter by continuously regenerating O2 that is consumed at the cathode, one must note that only minuscule amounts of species participate or are produced in these electrochemical reactions. Thus, the concentrations of H2O2 (before decomposition) and O2 in the 114 chamber media remain fairly constant for a period of time and are in equilibrium with that of the electrolyte. Because of this, stable baselines are obtainable prior to and after the addition of catalase solution into the chamber. Moreover, the response of the oxygen electrode to working H2O2 standards of various concentrations is found to be linear and proportional. Secondly, many common inhibitors (Switala et al., 2002) of catalase such as cyanide, azide, hydroxylamine, aminotriazole, and mercaptoethanol are absent in human urine samples. Urea, at concentrations above 4 M, was found to cause more than 50% denaturation of 1.12% catalase solution (Samejima et al., 1961). Although urea is a chief nitrogenous waste product excreted in urine, its average concentration in human urine is in the range of 100 to 600 mM (Gowrishankar et al., 1998; Kamel et al., 2004). Thus, urinary urea is not expected to significantly affect catalase performance, especially since each sample decomposition reaction itself takes not more than a minute. Furthermore, subjects had no restrictions on salt intake in their diet, so it is unlikely at any time in the studies that their urea excretions would reach or even exceed the upper limit of the abovementioned range (Gowrishankar et al., 1998). In 0.01M phosphate buffer at 20oC, catalase showed optimum activity at around pH 6.5 and little change in activity over the pH range of 5.5 to 7.5 (Bragger et al., 2000). Earlier studies have observed that the catalytic activity, spectra and sedimentation behaviour of catalase samples remain unaltered between pH 4.5 and 10, and even suggested a wider stability range of pH 3.5 to 11, outside of which true denaturation starts both for crystalline and lyophilized beef catalase samples (Tanford et al., 1962). In our studies, the pH of the urine samples lies within a narrow range of 5.9 to 7.1. So, 115 urinary pH is not likely to affect catalase performance significantly in the O2 electrode analyses. But how about urinary pH effects, if any, on the electrochemistry? Urine in the slightly acidic pH range would quench some of the OH- produced at the cathode halfreaction such that if the overall electrode reaction were to be reversible, Le Chatelier’s principle predicts that the equilibrium will shift to the right, leading to a greater rate of O2 consumption at the cathode and the system stabilizing at a higher baseline O2 level (due to a greater magnitude of deflection). However, this is not a reversible reaction, as the overall reaction thermodynamics is in favour of the formation of the more stable products (AgCl and OH-). Strictly speaking, it will be good to add buffer into the chamber to adjust the pH of the samples so as to eliminate any minuscule effects of pH. However, the use of smaller sample volumes to accommodate the buffer means compromising on the already poor sensitivity of the assay and inability to accurately measure H2O2, especially at concentrations of 5 µM and below. Nevertheless, it is important to point out here that in all urine spiking experiments with the O2 electrode, H2O2 added into urine samples is well-recovered in experimental calculation (80-120% recovery; Table 3.2) without any addition of buffers. These recovery data are supporting evidence that pH adjustment of urine samples may not be necessary. Heme-containing catalases degrade two molecules of H2O2 to one molecule of O2 and two molecules of H2O, involving the rapid interchange between the native, resting state porphyrin-iron (III) complex (ferricatalase) and the intermediate Compound I containing an oxoferryl group associated with a porphyrin π-cation radical (Chelikani et al., 2004). A study using a glucose oxidase-glucose system in buffer showed that when 116 the ratio of the rate of H2O2 generation (in µM per min) to the concentration of catalase (in µM) drops below 10 min-1, H2O2 is increasingly degraded through other catalytic pathways not leading to the liberation of O2 and involving other forms of catalase named Compounds II and III (de Groot et al., 2006). However, calculations show that the initial [H2O2]/[catalase] ratio at the point of introduction of catalase will always be above 10 in the chamber media (which contains 0.196 µM catalase) for all H2O2 working standards, the lowest one being 10 µM. In fact, based on this ratio requirement alone, urine samples with as low as 2 µM should be detectable. This shows that the unfavourable detection limit is largely determined by the limitations of the instrumental response and not due to the relative amounts of H2O2 and catalase present in the chamber. Unlike peroxidase, catalase is a lot more specific in its action. Ascorbate, up to a concentration of at least two times higher than that usually detected in human urine, did not affect catalase or the electrolytic reaction. In short, the O2 electrode assay is reliable for measuring urinary H2O2, based on the linear standard calibration plot, good recovery data and its almost non-existent susceptibility to interferences from samples. However, it unfortunately suffers from the disadvantage of lack of sensitivity and high detection limit. Non-enzymatic chemical-based methods Two non-enzymatic chemical-based assays studied in this project (the FOX-2 and FeTMPyP assays) were more sensitive to the detection of lower concentrations of H2O2 in water or PBS than the O2 electrode assay. Unfortunately, both assays suffered to different extents, from interfering compounds in urine such as ascorbate and urate. 117 The FOX-2 assay is actually not very specific for hydrogen peroxide because any sample oxidizing agent, irrespective of its chemical nature, can oxidize reagent ferrous to ferric ion which then can bind with xylenol orange to give the coloured complex with maximum absorption at 560nm (Banerjee et al., 2003). For instance, the FOX-2 assay can be made more specific for hydroperoxides by performing it in the presence and absence of triphenylphosphine, an agent which selectively reduces hydroperoxides to their corresponding alcohols and becomes triphenylphosphine oxide (Nourooz-Zadeh et al., 1994). In a similar way, the assay was made more specific for H2O2 by performing it in the presence and absence of catalase. How well the FOX-2 assay worked for a particular urine sample was dependant on the content of the urine, which in turn was influenced a lot by one’s diet. Ascorbate is known to be able to reduce Fe(III) formed by H2O2 back to Fe2+ (Halliwell, 1996), thus reducing the yield of Fe(III)-xylenol orange chromogen. For the same reason, the concentration of H2O2 in the aqueous humor determined by using FOX-1 assay was 5 to 12 times lower than by other assays; it was not surprising especially when ascorbate concentration in this ocular fluid was as high as 1.5 mM (Bleau et al., 1998). Sample dilution is not seen as an ideal solution to the problem of interferences as the sensitivity of the method goes down with higher number of dilutions. Peroxidase-based assays Peroxidase-based assays are worth examining because of their potential to offer even lower detection limits than non-enzymatic chemical-based assays. As low as 0.2 to 118 1 µM H2O2 in PBS could be detected when HVA, HPAA, amplex red, DCFH and DHR were used as the oxidizable substrates. However, most of these probes were later found to be unsuitable for urinary H2O2 analyses. For instance, the excitation/emission wavelengths employed for the detection of the dimerization products of HVA and HPAA were in the region where most of the fluorescence originated from the urinary matrix. Although the ABTS assay employed an absorbance wavelength that is close to the blue region of the visible spectrum where biological matrices are not likely to interfere spectrally, compounds with antioxidant activity against ABTS+• which were excreted in urine, rendered the assay useless. Peroxidase-based assays, although sensitive, generally lacked specificity. Amplex red, despite offering the advantage of high fluorescence power upon conversion to resorufin, was unable to compete effectively with other peroxidase substrates present in human urine for the HRP-compound I or II intermediates, to begin with. There was a limit to how much dilution of urine samples could help to remove the interferences affecting the amplex red assay. Among all the substrates studied, DCFH was the most reliable and promising. 2’,7’-Dichlorodihydrofluorescein (DCFH) assay Keston et al. (1965) originally described the use of DCFH as a useful specific indicator for H2O2 in the presence of peroxidase. It was already demonstrated that DCFH is oxidized by other species as well, such as ONOO- without peroxidase (Kooy et al., 1997), OH• (Myhre et al., 2003; Zhu et al., 1994), lipid hydroperoxides (Cathcart et al., 119 1983) and peroxidase in the absence of H2O2 (LeBel et al., 1992). ONOO- and lipid hydroperoxides are not expected in normal human urine, so they will not contribute as interferences to the DCFH assay. O2•-, however, could be generated in urine through reactions between autooxidizable compounds in urine with O2 from air after voiding (Halliwell et al., 2004). However, O2•- is not capable of oxidizing DCFH (Crow et al., 1997; LeBel et al., 1992; Zhu et al., 1994) and Myhre et al. (2003) have also classified DCFH as inappropriate for determining O2•-, HOCl and NO. Some other advantages of the DCFH assay include the probe’s high fluorescence power, sensitivity and possession of excitation/emission maxima (498/522 nm) which can sufficiently avoid interference from autofluorescence of urinary matrix. The DCFH reagent solution is stable for at least 24 hours, provided it is kept in the dark at 4oC. More importantly, neither catalase nor SOD (with or without H2O2) would oxidize DCFH to DCF (Rota et al., 1999a), so that catalase could be used freely for treatment of urine in control measurements while SOD could be used in the reaction progress study shown in Fig. 3.22. DCFH is usually commercially-available in the diacetate form (DCFH-DA) because of its prevalent use in cellular oxidative stress studies. DCFH-DA, being more hydrophobic, can diffuse easily across the cell membrane and be hydrolyzed by intracellular esterases to liberate DCFH which, upon reaction with oxidizing species forms its 2-electron oxidation product, the highly-fluorescent DCF (Bass et al., 1983). In our non-cellular work, base hydrolysis was performed to liberate DCFH for reaction with H2O2, in the presence of HRP (Fig. 4.1). 120 Fig. 4.1. Mechanism of DCFH-DA de-esterification to DCFH and further oxidation to highly-fluorescent DCF by ROS and RNS (adapted from Crow et al., 1997) Rota et al. (1999a) discovered that H2O2 oxidizes the resting state HRP-Fe(III) to HRP-compound I which will then react with DCFH to form the DCF semiquinone free radical (DCF•-) and HRP-compound II (Fig. 4.2). Other DCFH molecules can similarly reduce either newly-formed HRP-compound I or II to generate more DCF•-, but it is the reaction with compound II that regenerates the resting state HRP. Then DCF•- is airoxidized to DCF with the concomitant generation of O2•- (Rota et al., 1999a). In short, oxidation of DCFH to DCF by H2O2/peroxidase will form O2•- which upon disproportionation, spontaneously or by SOD, will generate more H2O2, so that the assay is inherently autocatalytic (Rota et al., 1999a). Moreover, photoreduction of DCF can occur in visible light, causing regeneration of DCF•- which can be oxidized by O2 once 121 again to produce more O2•- (Marchesi et al., 1999). Thus, experiments were conducted in as little light as possible to minimize the photoamplification effects. However, continuous fluorescence amplification of the HRP-catalyzed reaction was not observed in our studies, at least for the H2O2 working standards. Fig.3.21 shows that the fluorescence emission coming from each standard reaction mixture stabilized almost immediately for the lower H2O2 concentrations, while the highest standard concentration (10 µM) took not more than 10 minutes to stabilize. Incubation times of reaction were reduced from 30 minutes to 10 minutes. Even though the fluorescence of the sample reaction mixture went up with time, so did that of the corresponding catalase-treated sample reaction mixture, so that the differences between the 2 readings after 10 min and 30 min of incubation were not very different from each other, and gave similar urinary H2O2 concentration values. The H2O2-independent action of peroxidase with DCFH (LeBel et al., 1992) as well as the continuous generation of H2O2 in some urine samples under incubation (such that some H2O2 might react with DCFH/HRP instead of catalase) were most likely responsible for the rise in fluorescence with incubation time in the catalase-treated mixtures. Higher number of dilutions of urine would help to eliminate this rise (data not shown). 122 Fig. 4.2. Schematic representation of DCFH oxidation by HRP initiated by H2O2. DCF semiquinone free radical (DCF•-) reduces molecular oxygen to superoxide radical anion (a source of H2O2), consequently forming fluorescent DCF. H2O2, in turn, reacts with HRP, initiating another oxidative cycle (adapted from Bonini et al., 2006) Rota et al. (1999b) also reported that DCF can react with either HRP-compound I or II to give the DCF phenoxyl free radical (DCF•) and reduce the respective HRP enzyme intermediate. DCF• is structurally and chemically distinct from DCF•-. DCF• can oxidize many biochemical reducing agents in urine to free radicals, which may or may not, depending on their chemistry, react with O2 to form O2•- and ultimately, H2O2. For instance, DCF• reacts with ascorbate to give DCF and the ascorbate anion radical, but the ascorbate radical is relatively stable and does not further oxidize any other biochemical reductant in the system nor reduce any oxygen to O2•- (Rota et al., 1999b). So, a possible reason why some urine samples gave higher H2O2 values with the DCFH assay than the O2 electrode assay is that their urinary constituents promote a greater continuous generation of O2•- that ultimately amplifies the DCFH fluorescence, especially when there are many possible reactions (as we have discussed so far) involving the sample, DCFH, 123 DCF, air and/or light that can potentially lead to artificial radical formation. Meanwhile, autooxidation reactions in the corresponding catalase-treated sample reaction mixtures were greatly suppressed by the removal of H2O2 by catalase. The DCFH reaction progress study (Fig. 3.22) reinforces two important points: (a) The DCFH/HRP system reacts mainly with urinary H2O2, and not O2•-. SOD does not significantly inhibit the fluorescence emitted from the DCFH/HRP reaction. (b) The small reduction in fluorescence upon addition of SOD indicates that a small but significant fraction of H2O2 is possibly generated through autooxidation of compounds like polyphenolics in urine. The pO2 of urine within the bladder is considerably below that of ambient air and upon voiding, the urine is exposed to 21% oxygen so that O2 uptake occurs, leading to autooxidation and the production of O2•- which either spontaneously dismutates or causes further oxidation of other urinary constituents, eventually leading to the creation of more H2O2 (Long et al., 1999b; Halliwell et al., 2000a & 2004). Comparison study between the DCFH and the O2 electrode assays showed a lot of agreement (in about 85% of the tested urine samples). This demonstrated the suitability of this particular peroxidase-based assay in determining urinary H2O2 concentrations. Although neat urine samples could not be assayed accurately, not more than 20X dilutions of the samples were necessary to get consistent and reproducible results. As with other fluorogenic probes, DCFH may not be suitable for detecting H2O2 at concentrations higher than 10 or 20 µM due to limitations imposed by the nature of the method and reaction as well as the instrument used. An example is the substrate inhibition and reversible/irreversible inactivation of HRP by an excess amount of H2O2 at 124 the active site (Towne et al., 2004). However, linearity limitation is not a problem in our application since diluted urinary H2O2 values will always fall in between the linear range of 0.2 to 10 µM. Dilution of urine samples is necessary to reduce the matrix interference while at the same time increases the probability of reaction between H2O2 and DCFH/HRP. Dilution also helps to reduce the tendency of fluorescence to increase with time as a result of autooxidation reactions (data not shown). However, like any other peroxidase-based assays, the DCFH assay is also susceptible to interference coming from other peroxidase substrates present in urine, competing for the same active site on HRP while consuming H2O2. This may be why the assay reported lower urinary H2O2 concentrations than the O2 electrode for some urine samples. Ascorbate, however, could be less threatening as a competitor substrate in this assay; as mentioned earlier, it could quench DCF• produced from reaction of DCF with HRP enzyme intermediates back to DCF and become a very stable ascorbate anion radical (Rota et al., 1999b). On the whole, the DCFH assay is suitable for urinary H2O2 determinations based on its very low detection limit, high sensitivity, linearity of calibration plot, good agreement with the O2 electrode assay, excellent recoveries of spiked H2O2 in a variety of urine samples, good coefficient of variations in sample determinations (reproducibility) and high specificity for urinary H2O2. 125 Other previously-considered methods Some thought has been given to other fluorescent probes that are substrates of HRP. Resorufin (Brotea et al., 1988) and scopoletin (Fig. 4.3; Corbett, 1989) are probes that make use of inverse fluorescence measurements i.e. the starting material is fluorescent while the oxidized product is not. HRP-based assays using resorufin as a probe can be imagined as the exact inverse of the amplex red assay. Resorufin is highly fluorescent to start with and a low amount of H2O2 is going to be difficult to detect in such a high background. Resorufin assay is less sensitive and takes a long time to stabilize (Zhu et al., 1997). While naturally-occurring scopoletin may be reasonably stable in diffused light and not oxidized by either peroxidase or H2O2 alone (Corbett, 1989), it similarly lacks sensitivity. It has even lower fluorescent power than resorufin, has an excitation/emission short wavelength spectra (360/460 nm) that makes it susceptible to interference from autofluorescence of urinary matrix and may suffer interference from ascorbate (Gomes et al., 2005). Fig. 4.3. Chemical structure of scopoletin Serious consideration has been given to other methods which are non-peroxidase based but they too are deemed unsuitable for our purposes. Direct analyses of samples 126 with minimal clean-up and no derivatization by capillary electrophoresis sounds attractive but the detection limit is way too high, in the millimolar region (Shihabi et al., 2006). Determination of H2O2 using phosphine-based fluorescent reagents with sodium tungstate dihydrate (Onoda et al., 2006) or fluorescent probes based on a deprotection mechanism, similar to PFBSF (for example, Peroxyfluor-1; Chang et al., 2004), are not able to deliver sufficiently low detection limits, let alone selectivity, and they would require extensive organic synthesis. Purification or separation methods that make use of solid phase extraction cartridges to remove interfering or co-eluting compounds, and high performance liquid chromatography may result in losses or even artifactual generation of H2O2 through conversion and derivatization procedures that could be both timeconsuming and chemically harsh (Hamano et al., 1987). Basal urinary hydrogen peroxide measurements and coffee effect In short, a basal level of urinary H2O2 was found to be excreted by healthy subjects, though the basal level still varies from individual to individual due to the unique lifestyle and genetic make-up of each subject. Each subject experienced no drastic fluctuations in his/her creatinine-standardized urinary H2O2 throughout the day, especially with the DCFH assay. This assay also demonstrated less variation between the days for each subject than the O2 electrode and the effect of coffee consumption in raising urinary H2O2 concentration was proven in one subject. Very importantly, the potential suitability of urinary H2O2 as a biomarker of oxidative stress in the human body 127 was already demonstrated since the urinary H2O2 measured by the DCFH assay did not vary widely in the same subjects under the same conditions at different times. However, several authors (Kanabrocki et al., 2002; Pilger et al., 2002) reported marked daily intra-individual oscillations in some biomarkers of oxidative stress (including urinary 8OHdG, malondialdehyde and isoprostanes). Yuen et al. (2003) even concluded that the usefulness of urinary H2O2 as a potential biomarker for whole body oxidative stress was severely limited by wide intra- and inter-individual variations in concentration of H2O2, within and between days, unless the effects of disease or therapy induced very large changes in its concentration. On the other hand, Basu (2004) commented that there is no significant difference in urinary isoprostane levels in healthy subjects at any time of the day. Many a times, commonly-used assays are employed in research work to generate data, without much questioning. Perhaps, if the above authors except Basu (2003), had re-examined the reliability, suitability and accuracy of their chosen analytical methods, and employed a more robust assay with minimal interferences and high specificity, a different conclusion might have been reached. For instance, Yuen et al. (2003) obtained data with total reliance on a modified FOX assay that used a large sample volume (which would introduce more interfering species into the assay) and extended incubation time (giving ample time for the extraneous generation of H2O2 through auto-oxidation). Thus, the importance of using a good assay to generate data can never be overemphasized. 128 Possible future direction We have seen success with DCFH but failure with its structural analog, DHR, in the HRP-based assays. It would next be interesting to look at the suitability of dihydrofluorescein (HFLUOR; Hempel et al., 1999; Fig. 4.4) in our applications. HFLUOR is very structurally similar to DCFH, except for the absence of the 2’,7’dichloro subsituents. The outcome will enable us to infer the roles, if any, of the OH and Cl substitution, in the probes’ interactions with the HRP intermediate, which will decide their affinity for the enzyme relative to other peroxidase substrates present in urine. Ascorbate has an inhibitory effect to varying degrees in many of the peroxidasedependent assays we investigated (Martinello et al., 2006). A possible future experiment would be to see if commercially-available ascorbic acid quenchers (AAQs) in the form of 2,2,6,6-tetramethyl-1-piperidinyloxy (TEMPO) free radicals (Fig. 4.5) could be used to remove ascorbate in urine samples (Kayamori et al., 2000). The use of ascorbate oxidase could also be attempted. Urinary H2O2 can be studied alongside other classical biomarkers of oxidative stress in pathological conditions where oxidative stress is well-documented, as in malignancy (Banerjee et al., 2003) and in studies of the effect of intervention with antioxidants, health supplements or certain foods, often prescribed in therapy or diet, with the objective of controlling oxidative stress but are not routinely tested in clinical practice. Such intervention trials could in turn help to validate or disprove the biomarker concept. 129 Fig. 4.4. Comparison of structures of HFLUOR (dihydrofluorescein) and DCFH (2’,7’dichlorodihydrofluorescein), as well as their oxidized products (adapted from Hempel et al., 1999). Fig. 4.5. Molecular structures of ascorbic acid quenchers (AAQs; adapted from Kayamori et al., 2000) 130 CHAPTER 5 CONCLUSION Each single method has its own limitations and drawbacks, and therefore measurements of specific reactive species in biological systems or samples should always be made using at least two independent methods. It is beyond our scope to identify all possible compounds that are found in urine samples that can interfere with the specificity of these methods, though at least we know by now, that ascorbate, hydroquinones, nitrites and other phenolic compounds are substrates for peroxidases (Reszka et al., 2005) and these can be found in urine due to diet, oral medications, occupational exposures to chemical compounds and infections. Notwithstanding some of its limitations, the DCFH assay has proven to be reliable for measuring human urinary H2O2. Together with the O2 electrode assay, it can be used in future studies to further validate the suitability of urinary H2O2 as a biomarker of oxidative stress. For the first time, a relatively stable basal urinary H2O2 was measured in healthy individuals, and these values did not vary widely throughout the day, as well as between the days within a few months. Moreover, the effect of coffee consumption was easily detected by the assay. Thus, if we can get a good idea of an individual’s basal level of H2O2 excretion, we can use it to check for indication of onset/progression of disease (as clinical symptoms may appear too late), and investigate the effect of antioxidants therapy, health supplements and other foods for further validation of urinary H2O2 as a biomarker, by using a simple assay that can be carried out in any laboratory. 131 CHAPTER 6 REFERENCES Arnao, M.B., Cano, A., Hernandez-Ruiz, J., Garcia-Canovas, F. and Acosta M. (1996). 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Toxicol. 68, 582-587. 150 [...]... biomarkers of oxidative stress/ damage and the diseases with which they are associated 13 Table 1.2 Biomarkers of oxidative stress/ damage associated with some human diseases (adapted from Valko et al., 2007) NO2-Tyr, 3-nitrotyrosine 1.7 HYDROGEN PEROXIDE AS A BIOMARKER OF OXIDATIVE STRESS As mentioned earlier, H2O2 plays an important role as an inter- and intra-cellular signaling molecule, so a basal... The localization and effects of oxidative stress, as well as information regarding the nature of the ROS, may be gleaned from the analysis of discrete biomarkers of oxidative stress/ damage isolated from tissues and biological fluids Biomarkers are defined as characteristics that can be objectively measured and evaluated as indicators of normal biological processes, pathogenic processes, or pharmacologic... The data, as given in Table 1.3, show that for every collection, there was a significant intra-sample variation between the 3 assays The A2 2188 assay gave the lowest urinary H2O2 concentration values at all times while the O2 electrode assay gave the largest values Although the FOX-2 and O2 electrode assay gave values which 16 differed considerably in magnitude, a similar trend of increase and decrease... different samples were collected from him at the stated times (Table 1.3) within a day and were immediately analyzed by three assays, namely the oxygen electrode assay, the ferrous ion oxidation- xylenol orange version 2 (FOX-2) assay and the fluorescence assay The procedure used for the first two assays could be found in Chapter 2 The fluorescence assay was attempted using the amplex red-peroxidase assay. .. mitochondrial membrane and released into the matrix as well as the intermembranous space (Camello-Almaraz et al., 2006) O2•- is also produced from the direct reaction of autooxidizable molecules with dioxygen, as well as through the action of certain enzymes such as xanthine oxidase and galactose oxidase (Halliwell & Gutteridge, 1999) O2•- cannot directly attack DNA, proteins or lipids, but at elevated levels,... electrode assays) used in our laboratory and elsewhere; (b) develop a new assay suitable for the measurement of urinary H2O2 that is simple, accurate, sensitive, specific, reproducible and robust; and (c) use the assay developed in (b) to investigate if urinary H2O2 can meet as many of the requirements set out for an ideal biomarker of oxidative stress as possible 18 CHAPTER 2 EXPERIMENTAL PROCEDURES 2.1 MATERIALS... balance between their rates of production and their rates of removal by the antioxidant defence system which was briefly discussed earlier Oxidative stress occurs when there is a serious disturbance in this pro-oxidant – antioxidant balance in favour of the former, leading to potential damage (Sies, 1991) Oxidative stress can result from (Halliwell et al., 2004b): (a) Diminished levels of antioxidants,... Comparison of structures of HFLUOR (dihydrofluorescein) and DCFH (2’,7’-dichlorodihydrofluorescein), as well as their oxidized products 130 4.5 Molecular structures of ascorbic acid quenchers (AAQs) 130 xi LIST OF ABBREVIATIONS AND KEYWORDS AA Ascorbic acid AAQ Ascorbic acid quencher ABTS 2,2’-Azino-bis(3-ethylbenzothiazoline-6-sulfonic acid AscH- Ascorbate CuZnSOD Copper and zinc-containing superoxide... 3.9 Pentafluorobenzenesulfonyl fluorescein (PFBSF) 65 3.10 Oxidation of HVA in the presence of HRP to a fluorescence dimer 69 3.11 A standard calibration plot for the HVA assay 70 3.12 A standard calibration plot for the HPAA assay 74 3.13 Structure of ABTS and its oxidation products 76 3.14 A standard calibration plot for the ABTS assay 78 3.15 Chemical structures of polyphenols and their metabolites... to that of 8OHdG depends on the redox state of the cell and the presence of transition metal ions (Halliwell, 2000b) Hence, the same amount of free radical attack on DNA can give different levels of 8OHdG Another drawback is the artifactual generation of 8OHdG during DNA isolation from tissues, hydrolysis and analysis Consideration should also be given to other DNA base damage products which are known ... commonly-used biomarkers of oxidative stress/ damage and the diseases with which they are associated 13 Table 1.2 Biomarkers of oxidative stress/ damage associated with some human diseases (adapted from Valko... biomarkers of oxidative stress/ damage isolated from tissues and biological fluids Biomarkers are defined as characteristics that can be objectively measured and evaluated as indicators of normal biological... analyzed by DCFH assay 100 3.24 Effect of dilution of urine sample on DHR assay and comparison with DCFH assay 106 3.25 DHR assay recovery study and comparison with DCFH assay 107 3.26 Variations in