HEPATOCYTE MOLECULAR CYTOTOXIC MECHANISM STUDY OF AND ITS METABOLITES INVOLVED IN NONALCOHOLIC STEATOHEPATITIS AND

By

Yan (Cynthia) Feng

A thesis submitted in the conformity with the requirements

for the degree of Master of Science

Graduate Department of Pharmaceutical Sciences

University of Toronto

© Copyright by Yan (Cynthia) Feng 2010 ABSTRACT

HEPATOCYTE MOLECULAR CYTOTOXIC MECHANISM STUDY OF FRUCTOSE AND ITS METABOLITES INVOLVED IN NONALCOHOLIC STEATOHEPATITIS AND HYPEROXALURIA

Yan (Cynthia) Feng Master of Science, 2010 Department of Pharmaceutical Sciences University of Toronto

High chronic fructose consumption is linked to a nonalcoholic steatohepatitis (NASH) type of hepatotoxicity. is the major endpoint of fructose metabolism, which accumulates in the kidney causing renal stone disease. Both diseases are life-threatening if not treated. Our objective was to study the molecular cytotoxicity mechanisms of fructose and some of its metabolites in the liver. Fructose metabolites were incubated with primary rat hepatocytes, but cytotoxicity only occurred if the hepatocytes were exposed to non-toxic amounts of such as those released by activated immune cells. Glyoxal was most likely the endogenous toxin responsible for fructose induced toxicity formed via autoxidation of the fructose metabolite glycolaldehyde catalyzed by superoxide radicals, or oxidation by Fenton’s hydroxyl radicals. As for hyperoxaluria, glyoxylate was more cytotoxic than oxalate presumably because of the formation of condensation oxalomalate causing mitochondrial toxicity and oxidative stress. Oxalate toxicity likely involved pro-oxidant iron complex formation.

ii ACKNOWLEDGEMENTS

I would like to dedicate this thesis to my family. To my parents, thank you for the sacrifices you have made for me, thank you for always being there, loving me and supporting me throughout my life. To my uncle, thank you for providing me with many professional advices. And to Brandon, thank you for always believing in me, even in the times when I stopped believing in myself.

I am sincerely grateful to my supervisor, Dr. Peter J. O’Brien. Thank you for your constant guidance and support throughout my Master’s study at University of Toronto. Your enthusiasm and positive attitude for research have inspired me to overcome every challenge that I encountered during my study. Your insights and wisdom have always been helpful and led me to find the right path in my research.

I would like to give thanks to the past and current students in the lab. Thank you Rhea,

Qiang and Monica, for your friendship and company, and thank you for sharing your valuable knowledge and experiences with me. The current students, Luke, Kai, Stephanie and Sarah, thank you for all your helpful discussion and encouragement. I also want to thank the past members of the lab, for training me and helping me when I first started. All of you had made my experience extremely warm and memorable.

I would like to express my appreciation to my advisory committee members, Dr. Bruce and Dr. Pennefather, for their guidance and support. Finally I would like to acknowledge Dr.

Uetrecht and his lab for letting me use their plate reader.

iii TABLE OF CONTENTS

Abstract ii

Acknowledgements iii

Table of contents iv

List of publications and abstracts v

List of tables vi

List of figures vii

List of abbreviations viii-x

Chapter 1. General Introduction 1

Chapter 2. Materials and Methods 21

Chapter 3. Hepatocyte Inflammation Model for Cytotoxicity Research: 26 Fructose or Glycolaldehyde as a Source of Endogenous Toxins.

Chapter 4. Hepatocyte Study of the Molecular Cytotoxic Mechanisms of 40 Endogenous Toxins Formed by Hyperoxaluria

Chapter 5. Conclusions and Future Perspectives 67

References. 76

Appendix I. Permissions 87

iv LIST OF PUBLICATION AND ABSTRACTS

Publications:

Chan, K., Lehmler, H.J., Sivagnanam, M., Feng, C.Y., Robertson, L., O’Brien, P.J. (2009) Cytotoxic effects of polychlorinated biphenyl hydroquinone metabolites in rat hepatocytes. J Appl Toxicol. In press.

O’Brien, P.J., Feng, C.Y., Lee, O., Dong, Q., Mehta, R., Bruce, J., and Bruce, W.R. (2009). Fructose-derived endogenous toxins. Editors, O’Brien, P.J. and Bruce, W.R. Endogenous Toxins: Targets for Disease Treatment and Prevention. (pp173-204). WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim.

Feng, C.Y., Wong, S., Dong, Q., Bruce, J., Mehta, R., Bruce, W.R., and O’Brien, P.J. (2009). Hepatocyte inflammation model for cytotoxicity research: fructose or glycolaldehyde as a source of endogenous toxins. Arch. Physiol Biochem. 115(2), 105-111

Abstracts:

Feng, C.Y. and O’Brien, P.J. (2009) Rat hepatocyte molecular cytotoxic mechanism study of toxins formed by . Abstract presented at the National Health Research Day, University of Toronto, Toronto, Ontario. Nov 17, 2009.

O’Brien, P.J., Feng, C.Y., and Tafazoli, S. (2009). Accelerated cytotoxic mechanism screening (ACMS) for idiosyncratic drug hepatotoxin. Abstract presented at 12th Canadian Society for Pharmaceutical Sciences Annual Symposium, Toronto, Ontario. June 3-6, 2009.

Feng, C.Y. and O’Brien, P.J. (2009). Rat hepatocyte cytotoxicity model for primary hyperoxalurias. Abstract presented at the 6th Meeting of the Canadian Oxidative Stress Consortium, Winnipeg, Manitoba. May 7-10, 2009. (Runner-up for poster award)

v LIST OF TABLES

Table 1.1 Availability of high fructose corn syrup (HFCS) in the caloric sweetener supply in the United States.

Table 3.1 Fructose cytotoxicity is markedly enhanced by non-toxic H2O2/Fe(II) in isolated rat hepatocytes.

Table 3.2 The cytotoxicity of glycolaldehyde, a fructose metabolite, is markedly enhanced by non-toxic H2O2/Fe(II) in isolated rat hepatocytes.

Table 3.3 Glycolaldehyde protein carbonylation is increased by Fenton’s reagent and inhibited by a glyoxal scavenger in cell free system.

Table 4.1 Concentrations of each fructose metabolites studied that induce 50% cell death at 2 hours in isolated rat hepatocytes.

Table 4.2 Glyoxylate induced reactive species (ROS) formation in isolated rat hepatocytes.

Table 4.3 Endogenous glyoxylate reacts with mitochondrial oxaloacetate to form oxalomalate, a inhibitor.

Table 4.4 Concentrations of glycolate that induce 50% cell death at 2 hours in isolated rat hepatocytes under various stress conditions.

Table 4.5 Hydroxypyruvate induced ROS formation and lipid peroxidation in primary rat hepatocytes

Table 4.6 Oxalate induced ROS formation and lipid peroxidation in rat hepatocytes

vi LIST OF FIGURES

Figure 1.1 Simplified fructose to oxalate pathway and pathway.

Figure 1.2 The relationship between high fructose corn syrup, free fructose, total fructose intake and the prevalence of overweight and obese people in the United States.

Figure 1.3 Superoxide is detoxified by the superoxide dismutase, and can react with nitric oxide forming a more reactive oxidant peroxynitrite.

Figure 1.4 Detoxification of hydrogen peroxide could be catalyzed by catalase or reduced glutathione (GSH) peroxidise.

Figure 1.5 The Haber-Weiss reaction. Hydrogen peroxide is oxidized by transition metals by Fenton’s reaction forming highly reactive hydroxyl radicals.

Figure 1.6 Oxidation of guanine by hydroxyl radical forming 8-hydroxyguanine.

Figure 1.7 Schematic of advanced glycation end-product formation.

Figure 1.8 Schematic of lipid peroxidation formation.

Figure 1.9 Structures of some metabolites on the fructose to oxalate pathway.

Figure 3.1 The proposed molecular hepatotoxic mechanisms associated with fructose induced NASH as per the two-hit hypothesis.

Figure 4.1 Glyoxylate induced cytotoxicity in primary rat hepatocytes measured by the trypan blue exclusion test at 120min and 180min.

Figure 4.2 Hydroxypyruvate induced cytotoxicity in primary rat hepatocytes.

Figure 4.3 Oxalate induced cytotoxicity in primary rat hepatocytes.

Figure 4.4 Cysteine forms an adduct with glyoxylate.

Figure 4.5 Glyoxylate condenses with oxaloacetate forming oxalomalate, an inhibitor of aconitase and NADP+ dependent .

Figure 4.6 A simplified fructose to oxalate metabolic pathway.

vii LIST OF ABBREVIATIONS

ACMS Accelerated cytotoxic mechanism screening ADH AGT Alanine glyoxylate transaminase AGE Advanced glycation end-product ALDH ALE Advanced lipoxidation end-product ANOVA Analysis of variance ATP Adenosine-5-triphosphate BHA Butylated hydroxyl anisole BMI Body-mass index BSA Bovine serum albumin CBA 2-carboxybenzaldehyde Cl- Chloride CML Carboxymethyllysine

CO2 Carbon dioxide Cu Copper Cu(II) Cupric copper DETAPAC Diethylenetriaminepentaacetic acid DCFH-DA Dichlorofluorescein diacetate DCF Dichlorofluorescein DMSO Dimethylsulfoxide DNA Deoxyribonucleic acid DNPH 2,4-dinitrophenylhydrazine EDTA Ethylenediaminetetraacetic acid F.I. Fluorescent intensity Fe Iron

viii Fe(II) Ferrous iron Fe(III) Ferric iron FOX Ferrous oxidation in xylenol orange assay G/GO / glucose GI system Gastrointestinal system GLUT4 Glucose transporter 4 GLUT5 Glucose transporter 5 GR GSH Glutathione (reduced) GSSG Glutathione (oxidized)

H2O2 Hydrogen peroxide HCl Hydrochloric acid HFCS High fructose corn syrup HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid HPR Hydroxypyruvate reductase iNOS Inducible nitric oxide synthase JNK c-Jun N-terminal kinase pathway Concentration of the toxin lethal to 50% of the LC50 hepatocytes LDH MDA Malondialdehyde Na+ Sodium NAD+ Nicotinamide adenine dinucleotide (oxidized) NADH Nicotinamide adenine dinucleotide (reduced) NADP+ Nicotinamide adenine dinucleotide phosphate (oxidized) NADPH Nicotinamide adenine dinucleotide phosphate (reduced) NAFLD Nonalcoholic fatty liver disease NASH Nonalcoholic steatohepatitis OHQ 8-hydroxyquinoline

ix O2 Oxygen PH1 Primary hyperoxaluria type I PH2 Primary hyperoxaluria type II PPG Propargylglycine ROS Reactive oxygen species SOD Superoxide dismutase TBA Thiobarbituric acid TCA Trichloroacetic acid TNF-α Tumour necrosis factor alpha

x CHAPTER 1.

GENERAL INTRODUCTION

1.1. FRUCTOSE AND HEALTH CONCERNS

Fructose intake had been increased significantly in the past decades (Basciano et al.

2005). Major sources of dietary fructose include those from refined sugars such as

(50% fructose) and high fructose corn syrup (HFCS) (between 42%-90% fructose) present in many fast food and especially soft drinks. Form 1977 to 2001, total energy intake from soft drinks alone had increased form 3.9% to 9.2% (Nielsen and Popkin 2004).

HFCS was introduced in the 1970s, and has become more and more commonly used in the past decades. HFCS has been used to substitute for other caloric sweeteners in soft drinks and processed foods because it is much cheaper and sweeter when compared to an equal caloric portion of other sweeteners (Duffey and Popkin 2008). As shown in Table 1.1, HFCS use has increased from 0.5% of the total caloric sweetener in 1970 to 42% in 2004. Back in 1970,

HFCS 42 which contains 42% of fructose accounted for 100% of the HFCS use, whereas in

2004, about 60% came from HFCS 55 which contained 55% of fructose. Soft drinks and fruit drinks have been suggested to supply the most HFCS (Bray et al. 2004).

1 Table 1.1. Availability of HFCS in the caloric sweetener supply in the United States. Adapted from Bray et al. (2004)

Year HFCS Total caloric % HFCS of % HFCS 42* % HFCS (g/capita/day) sweeteners total caloric in total HFCS 55** in total (g/capita/day) sweeteners HFCS

1970 0.5 105.4 0.5 100 0 1975 4.3 100.7 4.3 100 0 1980 16.8 106.4 15.8 71.2 28.8 1985 40.0 111.6 35.8 34.3 65.7 1990 43.9 117.2 37.5 41.0 59.0 1995 51.0 127.5 40.0 39.9 60.0 2000 55.4 131.7 42.1 38.1 61.9 2004 52.4 124.8 42.0 40.5 59.5 Note: * HFCS 42 contains 42% of fructose. ** HFCS 55 contains 55% of fructose.

The increased chronic consumption of fructose had been associated with many health concerns. Fructose raises blood cholesterol and triglycerides, and increasing levels of these compounds contribute to cardiovascular disease. Fructose enriched diet could induce insulin resistance (Basciano et al. 2005), and a high intake of HFSC had been positively correlated with type 2 diabetes. This has been suggested to contribute to the pathogenesis of nonalcoholic fatty liver disease (NAFLD) (Collison et al. 2009; Ouyang et al. 2008). Research done previously by others suggested that a high fructose diet can induce changes in the c-Jun N-terminal kinase

(JNK), similar to that seen with inflammation (Kelley et al. 2004).

Fructose absorption and metabolism

Fructose absorption is distinctive from that of glucose. such as sucrose, enter the stomach, and are cleaved by to the glucose and fructose.

Glucose is absorbed by a sodium dependant transporter glucose transporter 4 (GLUT4) located on the apical membrane of intestinal cells. Fructose is absorbed in the duodenum and jejunum

2 of the intestine through the glucose transporter 5 (GLUT5). The absorption of fructose is by facilitated diffusion, therefore non-sodium dependent and non-energy requiring transporters are involved (Havel 2005). An increase in fructose intake in the diet was shown to increase GLUT5 expression in rats, which would lead to even more fructose being absorbed and metabolized

(Burant and Saxena 1994)

Glucose triggers insulin release from the beta cells of the pancreas, and insulin signals the insulin sensitive tissues (e.g. muscle cells, hepatocytes) to take up and store glucose, as well as stop the use of lipids as an energy source. Unlike glucose, fructose uptake and metabolism is not insulin dependent, as it is metabolized mainly by the liver. Fructose metabolism bypasses the rate limiting glycolysis regulatory step catalyzed by phosphofructokinase, and is phosphorylated to fructose-1-phosphate by fructokinase and adenosine triphosphate (ATP).

Following fructose infusion, a depletion of ATP and an accumulation of fructose-1-phosphate were observed (Bode et al. 1973). Chronic high consumption of fructose can up-regulate fructokinase mRNA, protein, and enzymatic activity in rats and human, leading to prolonged

ATP depletion (Ouyang et al. 2008).

Fructose and obesity

Fructose initiates lipogenesis in rats (Zavaroni et al. 1982) and humans (Bantle et al.

2000), and generates more triglycerides than any other (Hallfrisch 1990).

Fructose is rapidly metabolized catalyzed by fructokinase and ATP to form fructose-1- phosphate. Aldolase B then catalyzes the conversion of fructose-1-phosphate to glyceraldehyde and dihydroxyacetone phosphate which are intermediates in the glycolysis pathway. Glucose is

3 converted to glucose-6-phosphate catalyzed by glucokinase and ATP. Glucose-6-phosphate is then converted to fructose-6-phosphate catalyzed by isomerise, which is then converted to fructose-1,6-bisphosphate catalyzed by phosphofructokinase and ATP (Figure 1.1).

Phosphofructokinase is tightly regulated by allosteric inhibition by ATP and citrate (Havel

2005). Since fructose metabolism bypasses the phosphofructokinase regulatory step, it can continually supply intermediates for the glycolysis pathway. Furthermore, aldolase then catalyzes the conversion of fructose-1,6-bis phosphate to glyceraldehyde-3-phosphate and dihydroxyacetone phosphate. Dihydroxyacetone phosphate could be converted to glycerol-3- phosphate thus providing the glycerol portion of triglyceride synthesis (Havel 2005).

Glyceraldehyde-3-phosphate could proceed to generate pyruvate, which then forms acetyl-coA catalyzed by , thereby providing substrates for fatty acid synthesis. By supplying both glycerol-3-phosphate and the substrates for fatty acid synthesis, fructose triggers the de novo synthesis of triglycerides (Mayes 1993). Observations that glycerol-3-phosphate and pyruvate level increases following fructose injection intravenously (Zakim and Herman

1968) and intraperitoneally (Burch et al. 1969) further supports the fact that fructose contributes to lipogenesis by providing both the glycerol and acetyl component.

4 Fructose Glucose Glucokinase Fructokinase Glucose-6-P Fructose-1-P

Aldolase B Fructose-6-P

Dihydroxyacetone-P Glyceraldehyde Phosphofructokinase

Fructose-1,6-Bis-P

Glycerol-3-P D-glycerate

Glyceraldehyde-3-P Dihydroxyacetone-P Hydroxypyruvate Glyceraldehyde-3-P ROS dehydrogenase Methylglyoxal ROS 1,3-bis-P-glycerate Glyoxal Glycolaldehyde

Glycolic acid

Pyruvate

Glyoxylic acid

Acetyl CoA

Glycine Oxalic acid

Figure 1.1. Simplified fructose to oxalate pathway and glycolysis pathway. Fructose metabolism bypasses the glycolysis regulatory enzyme phosphofructokinase, resulting in a continual supply of the substrates to the glycolysis pathway, contributing to lipogenesis by providing both the glycerol-3-phosphate and acetyl coA. Modified from Danpure and Rumsby 2004).

Unlike glucose, fructose intake does not stimulate insulin or leptin secretion (Bray et al.

2004). Following glucose intake, insulin secretion by beta cells in the pancreas is induced to

increase the glucose transporter on the cell surface, and to stimulate leptin secretion which

promotes the feeling of satiety (Saad et al. 1998). Pancreatic beta cells do not express fructose

transporter GLUT 5, therefore fructose does not stimulate insulin release or the associated

responses (Sato et al. 1996). The effect of the absence of leptin is best demonstrated in leptin

5 deficient individuals who are usually obese, and whose low leptin levels may prolong hunger and desire for food, resulting in an increase in body fat (Bray et al. 2004). Overall, as shown in

Figure 1.2, fructose intake, and in particular HFCS intake is closely linked to the prevalence of obese and overweight individuals in United States (Bray et al. 2004).

Figure 1.2. The relationship between intake of HFCS (♦), free fructose (▲), total fructose intake(●) and the prevalence of overweight (■) and obese people (x) in United States (Bray et al. 2004).

1.2. NAFLD AND NASH

Nonalcoholic fatty liver disease (NAFLD) describes a range of conditions including steatosis (fatty liver), nonalcoholic steatohepatitis (NASH), and more severe hepatic fibrosis and cirrhosis (irreversible scarring of the liver tissue) (Browning and Horton 2004). All of these conditions are characterized by fatty infiltration of hepatocytes in the absence of alcohol use.

6 NAFLD is closely associated with obesity; the population with a body-mass index (BMI) of more than 30 have a 4.6 times higher prevalence of NAFLD compared to the population with a normal BMI (Cavicchi et al. 2000). NAFLD is also closely correlated with insulin resistance, type II diabetes and hyperlipidemia (Angulo and Lindor 2002). It is most commonly present in developed countries where people are exposed to a high calorie, high sugar, and high fat diet, as well as a sedentary life style. In the United States, the prevalence of NAFLD is higher than the prevalence of alcoholic fatty liver disease and chronic viral liver disease combined, suggesting

NAFLD is the leading cause of liver dysfunction (Clark et al. 2003). It was proposed that steatosis (fatty liver) affects more than two thirds of the obese population (BMI 30 – 40), and over 90% of the morbidly obese (BMI > 40) population, whereas steatohepatitis (fatty liver with inflammation) affects 3% of normal individuals, 19% of the obese population, and more than 50% of morbidly obese individuals (Angulo and Lindor 2002).

NASH describes a more serious condition of NAFLD associated with inflammation and fibrosis besides fatty infiltration in the hepatocytes. It can progress to cirrhosis, liver failure, and hepatocellular carcinoma. NASH is the leading cause of liver dysfunction and cirrhosis in nonalcoholics in Europe and America (Angulo and Lindor 2002). Not all steatosis progresses to

NASH, and a two-hit hypothesis had been proposed (Day and James 1998). The first hit is steatosis and insulin resistance, and the second hit is oxidative stress coming from inflammation, xenobiotics or drugs. In the absence of the second hit, only simple steatosis would take place, and NASH would not ensue.

7 1.3. HYPEROXALURIA

Hyperoxaluria is characterized by elevated endogenous oxalate, which readily crystallizes with calcium forming insoluble salts. Oxalate is present in the diet and absorption of free oxalate anion occurs in the gastrointestinal system through passive, non-energy required diffusion (Binder 1974). However calcium inhibits oxalate uptake. The majority of oxalate is synthesized from dietary sources especially fructose and sucrose. Since the primary route of endogenous oxalate excretion is through the urinary system, calcium oxalate could cause stone formation in the kidney, renal tubules, liver, brain, and bones (Friedman et al. 1962; Williams

1978).

There are four categories of hyperoxaluria. (1) Primary hyperoxaluria is due to genetic defects in the detoxification of glyoxylate, an oxalate precursor. (2) Enteric hyperoxaluria is caused by increased absorption of oxalate due to inadequate calcium binding to oxalate in the gastrointestinal system, and is usually associated with chronic diarrhea. (3)

Dietary hyperoxaluria results from an increased intake of food high in oxalate content, which includes e.g. spinach (970mg/g), rhubarb (805mg/g), and cocoa (700mg/g) (Noonan and Savage

1999). (4) Finally, idiopathic or mild hyperoxaluria, the most common type of hyperoxaluria, may be caused by increased dietary intake of oxalate or elevated endogenous production.

Furthermore, dietary fructose consumption is also closely linked to hyperoxaluria. Among glucose and some other sugar alcohols, fructose produces the most oxalate (Rofe et al. 1980).

Following fructose infusion, increases in both urinary oxalate and calcium were observed

(Nguyen et al. 1995).

8 Primary hyperoxaluria is a rare autosomal recessive genetic disorder, it is a severe form of hyperoxaluria caused by enzymatic deficiency in the liver. In primary hyperoxaluria, the detoxification enzymes of glyoxylate are deficient, resulting in an increased oxalate formation from glyoxylate. There are two types of primary hyperoxaluria with distinctive enzyme deficiencies. Primary hyperoxaluria type I (PH1) is caused by defective alanine glyoxylate transaminase (AGT), and will result in increased production of oxalate and glycolate from glyoxylate. Primary hyperoxaluria type II (PH2) results from defective glyoxylate reductase

(GR), also known as hydroxypyruvate reductase (HPR), and will result in increased oxalate production from glyoxylate and L-glycerate production from hydroxypyruvate. Primary hyperoxaluria is associated with elevated oxidative stress, such as increased free radical formation (Scheid et al. 1996; Thamilselvan et al. 2000), lipid peroxidation (Thamilselvan et al.

2000) and mitochondrial damage (Cao et al. 2004).

1.4. OXIDATIVE STRESS

Reactive oxygen species (ROS) are produced as a natural by-product of normal metabolism of oxygen. For example, they are generated by NADPH oxidase in neutrophils when they are activated and are involved in the killing of foreign bacteria and pathogens. They have important roles in cellular signalling toward cell migration, gene expression, and cell growth, and are essential for cellular function. On the other hand, ROS could create damage to

DNA, protein, and cell membrane. Cells possess antioxidant defence such as antioxidant enzymes and small molecules that can protect against ROS induced cellular damages. There’s normally a well-maintained balance between level of pro-oxidant and antioxidant. When the

9 levels of pro-oxidants increase to the degree that they cannot be detoxified fast enough, there will be an imbalance between pro-oxidants and anti-oxidants, and oxidative stress will take place. Oxidative stress is believed to be linked to the pathogenesis of many diseases including neurodegenerative diseases, cardiovascular diseases, and cancers. Some of the ROS molecules

1 causing liver injury include superoxide (O2·-), singlet oxygen ( O2), hydrogen peroxide (H2O2), and hydroxyl radicals (·OH) (Browning and Horton 2004).

Superoxide radical

Superoxide radical (O2·-) forms when O2 undergoes one electron reduction. Superoxide radicals can be produced by reduced nicotinamide adenine dinucleotide phosphate (NADPH) oxidase or , but it is generally agreed that the major site of superoxide production is in the mitochondria, through electron leakage from the electron transport chain at complex I and complex III (Turrens 1997). The fructose oxidation product glyoxal has been shown to inhibit the respiratory chain and therefore increases superoxide production (Rosca et al. 2005). Superoxide is detoxified enzymatically by superoxide dismutase (SOD) forming hydrogen peroxide (Figure1.3) (McCord and Fridovich 1969). SOD is present in the mitochondria and cytosol where superoxide is produced. Furthermore, superoxide reacts with nitric oxide radical (·NO) rapidly and non-enzymatically forming a powerful oxidant peroxynitrite (ONOO-) (Pacher et al. 2007) (Figure 1.3). It was suggested by Beckman and

Koppenol that the nitric oxide radical was the only biological molecule produced in a quantity high enough that it would react fast enough to out-compete SOD and react with superoxide.

The product peroxynitrite is more reactive than either of its parent molecules (Beckman and

10 Koppenol 1996). Peroxynitrite could directly oxidize DNA and proteins, or indirectly cause damage via radical mediated mechanisms (Pacher et al. 2007).

Superoxide Dismutase (1) O2 ·- O2 + H2O2

2 H+

O ·- + ·NO ONOO- (2) 2

Figure 1.3. Superoxide is detoxified by the enzyme superoxide dismutase forming oxygen and hydrogen peroxide (1). Superoxide can react with nitric oxide forming a more reactive oxidant peroxynitrite (2) that can directly damage DNA and protein.

Hydrogen peroxide

Peroxisomes are important in the metabolism of various substrates, such as fatty acid, D- amino acid, and purines (Angermuller et al. 2009). It contains various crucial to its function, by removing hydrogen from the and transferring it to an O2 acceptor, thereby producing hydrogen peroxide (H2O2). were suggested to be responsible for about 35% of the total hepatic H2O2 (Boveris et al. 1972). H2O2 can oxidize protein thiol groups, and undergo Fenton’s reaction with transition metals, generating the much more reactive hydroxyl radical (Halliwell and Gutteridge 1992). H2O2 is detoxified by peroxisomal catalase forming water and oxygen (Figure 1.4). H2O2 is also detoxified by glutathione with reduced glutathione (GSH) as a (Figure 1.4) (Tappel et al. 1982). In the process, GSH is oxidized to oxidized glutathione (GSSG,) and requires glutathione reductase and NADPH to be reduced back to GSH.

11 Catalase 2 H2O2 2 H2O + O2 (1)

GSH peroxidase H2O2 2 H2O (2)

2GSH GSSG

Figure 1.4. Detoxification of H2O2 could be catalyzed by catalase forming water and oxygen (1), or by GSH peroxidase using GSH as a cofactor forming water and GSSG (2).

Iron and Fenton’s reaction

Iron (Fe) is an essential transition metal for maintaining cellular enzymatic and protein functions. Iron is tightly regulated by binding to ferritin, and the liver is the main site for iron storage. In the presence of transition metals such as Fe and copper (Cu), H2O2 undergoes the

Haber-Weiss reaction (Fenton’s reaction), generating the highly reactive oxidant, hydroxyl radical (·OH) (Figure 1.5) (Chance et al. 1973).

3+ 2+ Fe + O2·- Fe + O2

2+ 3+ - Fenton’s reaction Fe + H2O2 Fe + ·OH + OH

- Net reaction O2·- + H2O2 O2 + OH + ·OH

Figure 1.5. The Haber-Weiss reaction. H2O2 is oxidized by transition metals by Fenton’s reaction forming highly reactive hydroxyl radicals.

Hydroxyl radicals have been suggested to be the most powerful ROS radical. It can oxidize membrane polyunsaturated fatty acid causing lipid peroxidation. It also directly modifies protein amino acid groups and purine and pyrimidine bases of DNA (Breen and

Murphy 1995) causing mutations. One example of such DNA modification is the reaction of hydroxyl radical with guanine forming 8-hydroxyguanine, which is widely used as the marker

12 of oxidative DNA damage (Figure 1.6) (Valko et al. 2006). There is no enzyme that could directly catalyze the detoxification of hydroxyl radical, as the radical is very reactive and reacts with most biomolecules.

O O N N HN HN + ·OH OH N N N N H2N H2N H H

Figure 1.6. Oxidation of guanine by hydroxyl radical forming 8-hydroxyguanine, a commonly assayed marker for oxidative DNA damage (Valko et al. 2006).

1.5. ADVANCED GLYCATION AND LIPOXIDATION END-PRODUCTS

Advanced glycation end-product (AGE)

Long term glycemia could result in formation of advanced glycation end-products

(AGE). Carbonyl groups of sugars and amino groups of protein, especially lysine and arginine, can undergo a non-enzymatic Maillard reaction forming a Schiff base. Schiff base formation is usually reversible, but can go through rearrangements forming a stable Amadori product. After more oxidative and non-oxidative reactions, irreversible advanced glycation end-products

(AGE) are formed (Figure 1.7) (O'Brien et al. 2005). One of the most well-known AGE products is carboxymethyllysine (CML), that is implicated in many disease such as diabetes and renal failure, and is used generally as a marker of oxidative damage (Kislinger et al. 1999).

AGE could form cross-links between proteins altering their structure and function. AGE formation contributes to the progression of insulin resistance (Unoki and Yamagishi 2008) and atherosclerosis (Vlassara et al. 1995), and increased AGE levels been found to be associated

13 with aging, diabetic complications (McCance et al. 1993), NASH (Hyogo et al. 2007) and cardiovascular disease (Peppa and Raptis 2008). AGE, especially glyceraldehyde derived AGE, was found to be significantly elevated in NASH patients (Hyogo et al. 2007). Sugars differ in their ability to react with proteins, and fructose was shown to be more reactive than glucose and thus forms AGE more readily compared to glucose (Suarez et al. 1989).

Advanced lipid peroxidation product (ALE)

Free radicals (e.g. hydroxyl radicals) can oxidatively modify polyunsaturated fatty acids causing lipid peroxidation. Fatty acid undergoes oxidation forming a fatty acid radical, which reacts with oxygen forming a lipid peroxyl radical. This then reacts with another fatty acid, generating a lipid peroxide and a new lipid radical. The reaction propagates forming lipid peroxidation products such as malondialdehyde (MDA) (Figure 1.8) (Young and McEneny

2001). The chain reaction stops only if two radicals meet, or when an anti-oxidant donates its electron. Lipid peroxidation products could irreversibly modify proteins forming advanced lipoxidation products (ALE). MDA, for example, can modify protein lysine groups forming

MDA-lysine adducts, implicated in atherogenesis and is used as a marker for oxidative stress

(Huang et al. 1999). ALE can cause protein cross-links, modify lipoproteins, and alter protein enzyme activities. Similar to AGE, elevated ALE levels are implicated in cardiovascular and diabetic complications (Shanmugam et al. 2008).

14

Figure 1.7. Schematic of advanced glycation end-product (AGE) formation. reacts non-enzymatically with protein forming a reversible Schiff base. Schiff base undergoes rearrangement forming a stable Amadori product, which go through more rearrangements forming AGE. (O'Brien et al. 2005)

15

Polyunsaturated lipid

+ OH

Lipid radical

Rearranged

O2

Lipid peroxyl radical

O O +

Another polyunsaturated lipid

Lipid peroxide Malondialdehyde 4-hydroxynonenal O etc. OH +

Lipid radical

Figure 1.8. Schematic of lipid peroxidation formation. Radicals, such as a hydroxyl radical, oxidatively modify polyunsaturated fatty acid forming a lipid radical. Lipid radical reacts with oxygen forming a peroxyl radical, which then reacts with another fatty acid forming a lipid peroxide and a new lipid radical. This reaction propagates forming lipid peroxidation product such as malondialdehyde (MDA) which could irreversibly modify proteins forming advanced lipoxidation product (ALE). Modified from Young and McEneny (2001).

16 1.6. SCOPE OF THESIS

Fructose intake is implicated in both NASH (Collison et al. 2009; Ouyang et al. 2008) and hyperoxaluria (Rofe et al. 1980). The molecular cytotoxic mechanisms for both conditions are still elusive, and requires a good model of study. Since fructose metabolism and oxalate production occur mainly in the liver, our study in primary rat hepatocyte should serve as a first step in modelling the conditions leading to toxicity of fructose and its metabolites. In Chapter 3, the hepatocyte inflammation mode (Tafazoli et al. 2008 ; Tafazoli and O’Brien 2009) was used to study the toxicities of fructose and one of its most genotoxic metabolites, glycolaldehyde. In

Chapter 4, fructose metabolites involved in hyperoxaluria and the metabolism of the intermediates hydroxypyruvate, glycolate, glyoxylate and oxalate, were studied in the hepatocytes using “Accelerated Cytotoxic Mechanism Screening (ACMS )” techniques (Chan and O'Brien 2006; O'Brien and Siraki 2005). Structures of some fructose metabolites are shown in Figure 1.9.

17

Fructose HOH2C O OH HO

CH2OH OH

Glyceraldehyde O O HO

Dihydroxyacetone O HO OH

Glyceric acid O

HO OH

OH

Hydroxypyruvic acid HO O

OH O Glycolaldehyde OH O

Glycolic acid O OH

OH Glyoxylic acid O OH

O Oxalic acid O OH

HO O Figure 1.9. Structures of some metabolites on the fructose to oxalate pathway.

18 1.7. HYPOTHESES

Background to Hypothesis 1 (Chapter 3)

Fructose and glycolaldehyde are both toxic under simulated conditions of inflammation

(H2O2 formation by inflammatory or immune cells). ACMS techniques showed that the molecular cytotoxic mechanism involved oxidation of fructose and glycolaldehyde by hydroxyl radicals to form carbon centered radical intermediates such as glycolaldehyde and glyoxal radicals to form glyoxal. Glyoxal markedly increased hepatocyte susceptibility to H2O2 by at least a hundred fold and also increased endogenous H2O2 production thereby causing oxidative stress.

Hypothesis 1

“H2O2/Fe(II) (Fenton products) in hepatocytes oxidize fructose and its metabolite glycolaldehyde intracellularly to form glyoxal which markedly increases oxidative stress cytotoxicity”

Background to Hypothesis 2 (Chapter 4)

Glyoxylate, but not calcium oxalate, is a major endogenous toxin in hyperoxaluria.

Glyoxylate induced toxicity involves its carbonyl group, as well as formation of condensation products with oxaloacetate causing mitochondrial toxicity. Oxalate induced cytotoxicity involves pro-oxidant complex formation with transition metals.

Hypothesis 2 a) Glyoxylate is more effective than oxalate at causing mitochondrial toxicity and cytotoxicity. b) Iron oxalate is more effective than calcium oxalate at causing oxidative stress cytotoxicity.

19 1.8. ORGANIZATION OF THESIS

This thesis is organized in the manner of a traditional thesis with sections of Introduction,

Material and Methods, and Discussion. Chapter 3 of the thesis is a paper written with me as first author that was published in the journal “Archives of Physiology and ” (Feng et al. 2009). Chapter 4 is a paper about to be submitted to a journal for publishing. Material and methods in Chapter 2 apply to both Chapter 3 and Chapter 4.

20 CHAPTER 2

MATERIALS AND METHODS

2.1. CHEMICALS

Type II collagenase was purchased from Worthington (Lakewood, NJ). Fructose was purchased from Alfa Aesar. Bovine serum albumin (pH 7, ≥98%, Sigma product number

A7906) and all other chemicals were purchased from Sigma-Aldrich Co. (Oakville, ON).

2.2. ANIMAL TREATMENT

Male Sprague-Dawley rats weighing 275-300 grams were purchased from Charles River

Laboratories. All animal care and experimental procedures were carried out according to the guidelines of the Canadian Council on Animal Care (CCAC 1993). Rats were housed in ventilated plastic cages over PWI 8-16 hardwood bedding. There were 12 air changes per hour,

12 hour light photoperiod (lights on at 0800 h) and an environmental temperature of 21-23 °C with a 50-60% relative humidity (Moldeus et al. 1978). The animals were fed with a normal standard chow diet and water ad libitum.

2.3. HEPATOCYTE PREPARATION

Hepatocytes were isolated from rats by collagenase perfusion of the liver as described by Moldeus and coworkers (Moldeus et al. 1978). Isolated hepatocytes (106 cells/ml, 10 ml) were suspended in Krebs-Henseleit buffer (pH 7.4) containing 12.5mM 4-(2-hydroxyethyl)-1- piperazineethanesulfonic acid (HEPES) in 50ml round-bottomed flasks continually rotating in a

37 °C water bath, under an atmosphere of 95% O2 and 5% CO2. Hepatocytes were allowed to

21 acclimatize for 30 min before the addition of chemicals. Stock solutions of chemicals were prepared in Millipore water or dimethylsulfoxide (DMSO), immediately prior to use.

2.4. HEPATOCYTE VIABILITY

Hepatocyte viability was assessed microscopically by detecting plasma membrane disruption determined by the trypan blue (0.1%, w/v) exclusion test (Moldeus et al. 1978) at 30,

60, 120, and 180 min during a 3 hour incubation period. The cells were at least 80-90% viable before use. Catalase inhibited hepatocytes were prepared by preincubating hepatocytes with

4mM sodium azide for 10 min (Silva and O’Brien 1989). Glutathione (GSH) depleted hepatocytes were prepared by preincubating hepatocytes with 200μM 1-bromoheptane for 30 min (Khan and O'Brien 1991). AGT inhibited hepatocytes were prepared by preincubating hepatocytes with 5mM propargylglycine (PPG) for 30 min (Cornell et al. 1984). GR/HPR inhibited hepatocytes were prepared by addition of 10mM 2-carboxybenzaldehyde (CBA)

(Litvinovich and O'Brien 2007).

2.5. HEPATOCYTE INFLAMMATION MODEL

Hepatocytes susceptibility to hepatotoxin was found to be markedly increased when exposed to the products of activated immune cells such as H2O2 generated by their activated

NADPH oxidase (Tafazoli et al. 2008; Tafazoli and O’Brien 2008). However, following the addition of H2O2 to the hepatocytes, H2O2 was metabolized by catalase within a minute (Ou and

Wolff 1996). Therefore, a H2O2 generating system consisting of glucose and glucose oxidase was used to avoid rapid metabolism of H2O2. This system was shown to continuously supply

H2O2 without affecting the hepatocyte viability or GSH levels (Tafazoli et al. 2008).

22 2.6. PROTEIN CARBONYLATION ASSAY IN ISOLATED RAT HEPATOCYTES

The total protein bound carbonyl content was assessed by derivatizing the protein carbonyl adducts with 2,4-dinitrophenylhydrazine (DNPH). Briefly, 0.5 ml of hepatocyte

(0.5 × 106 cells) was added to 0.5 ml of DNPH (0.1%, w/v) in 2 N hydrochloric acid (HCl) and incubated at room temperature in dark for 1 h. The reaction was terminated by the addition of

1ml of trichloroacetic acid (TCA) (20%, w/v). Centrifugation at 50×g pelleted the protein, and the supernatant was removed. Excess unbound DNPH was extracted by washing with 0.5 ml of ethyl acetate:ethanol (1:1) solution for three times. The remaining pellet was dried under a stream of nitrogen and dissolved in 1 ml of Tris-buffered 8.0 M guanidine–HCl (pH 7.2). The solubilized hydrazones were measured spectrophotometrically at 370 nm (Hartley et al. 1997).

2.7. DETERMINATION OF PROTEIN CARBONYL CONTENT OF BOVINE SERUM

ALBUMIN (BSA)

In a cell free assay, the total protein bound carbonyl content of bovine serum albumin

(BSA) was determined by derivatizing the protein carbonyl adducts with 2,4- dinitrophenylhydrazine (DNPH). BSA (2 mg/ml) was prepared in 100 mM phosphate buffer

(pH 7.4) Briefly, BSA (0.5 ml) was incubated for 1 h at room temperature with 0.5 ml of

DNPH (0.1% w/v) in 2 N HCL. 1 ml of TCA (20% w/v) was added to the suspension to stop the reaction. The sample was centrifuged at 50×g to obtain the cellular pellet, and the supernatant was removed. DNPH was removed by extracting the pellet three times using 0.5 ml of ethyl acetate:ethanol (1:1) solution. After the extraction, the pellet was dried under a gentle stream of nitrogen and dissolved in 1 ml of Tris-buffered 8.0 M guanidine-HCl (pH 7.2). The solubilized hydrazones were measured at 370 nm (Hartley et al. 1997).

23 2.8. HEPATIC HYDROGEN PEROXIDE FORMATION

Hepatic hydrogen peroxide (H2O2) was measured by reacting with FOX reagent (ferrous oxidation in xylenol orange assay). Briefly, 50 μl of the hepatocytes suspension

(0.05 × 106 cells) was added to 950 μl of FOX reagent and incubated at room temperature in the dark for 30 min. The FOX reagent consisted of 25 mM sulfuric acid, 250 μM ferrous ammonium sulfate, 100 μM xylenol orange and 0.1 M sorbitol. Following incubation, samples were analyzed spectrophotometrically at 560 nm. The extinction coefficient

5 −1 −1 2.35 × 10 M cm was used to quantify the molar concentration of H2O2 through Beer's law

(Ou and Wolff 1996).

2.9. HEPATIC LIPID PEROXIDATION

Lipid peroxidation products, such as malondialdehyde (MDA), react with thiobarbituric acid (TBA) forming a pink adduct that can be measured spectrometrically. Briefly, 1 ml aliquots of hepatocytes from each sample were taken out at various time points after treatment.

250µl of trichloroacetic acid (TCA) (70% w/v) was added to lyse the cells, followed by addition of 750µl of water and 1 ml of TBA (0.8%, w/v). Suspensions were placed in boiling water for 20 min, and centrifuged at a high speed for 5 min after cooling. Supernatant was read spectrophotometrically at 532nm (Smith et al. 1982).

2.10. HEPATIC REACTIVE OXYGEN SPECIES (ROS) FORMATION

Hepatic ROS formation was determined using 2,7-dichlorofluorescein diacetate (DCFH-

DA). DCFH-DA enters hepatocytes, and is hydrolyzed to non-fluorescent dichlorofluorescein, which then reacts with ROS forming highly fluorescent dichlorofluorescein (DCF) and effluxes

24 the cell. 1 ml aliquots of hepatocyte from each sample were taken out at various time points after treatment, and were centrifuged for 1 min at 50×g. Following centrifugation, the supernatant was discarded, and the pellet was resuspended in 1 ml of Krebs-Henseleit buffer containing 1.6 μM of DCFH-DA. After 10 min incubation in dark at 37°C with gentle shaking, the fluorescent intensity of ROS product was measured at 490nm excitation and 520nm emission wavelengths (Shangari and O'Brien 2004).

2.11. HEPATIC MITOCHONDRIAL MEMBRANE POTENTIAL

Hepatic mitochondrial membrane potential was determined by the uptake of cationic fluorescent dye, rhodamine 123. Briefly, 500µl aliquots of hepatocytes from each sample were taken out at various timepoints after treatment, and centrifuged at 50×g for 1 min. The supernatant was discarded, and pellet was resuspended in 2 ml of Krebs-Henseleit buffer containing 1.5 uM rhodamine 123, and incubated for 10 min in dark at 37°C with gentle shaking. Hepatocytes were then separated by centrifugation, and the amount of rhodamine 123 remained in the incubation medium was measured fluorimetrically at 490nm excitation and

520nm emission wavelengths (Andersson et al. 1987).

2.12. STATISTICAL ANALYSIS

Statistical analysis was performed by a one-way ANOVA test. Significance between treatments was assessed by employing Tukey’s post hoc test. Results are presented as mean ± standard error (S.E.) for three separate experiments. Values were considered to be significant if p<0.05. SPSS version 16.0 for Windows was used to perform the statistical analysis.

25 CHAPTER 3

HEPATOCYTE INFLAMMATION MODEL FOR CYTOTOXICITY RESEARCH: FRUCTOSE OR GLYCOLALDEHYDE AS A SOURCE OF ENDOGENOUS TOXINS

3.1. INTRODUCTION

The high chronic consumption of fructose is associated with health concerns. An increase in obesity in the population over the past 35 years has coincided with an increase in the consumption of sugars (22% increase) and fats (48%) by the population. Fructose intake has also increased partly as a result of high-fructose corn syrup (HFCS)-55 ingestion, and unlike glucose ingestion, sucrose or fructose enriched diets have caused a sustained elevation of plasma triglycerides in the human population, which suggested that chronic over consumption of fructose could contribute to atherogenesis and cardiovascular disease (Stanhope et al. 2008).

Sucrose- and fructose-enriched diets also caused hepatic insulin resistance in rats independently of obesity that involved c-Jun N-terminal kinase (JNK) activation (Kelley et al., 2004). Glucose is metabolized by most cells whereas fructose is mostly metabolized in vivo by the liver, and now 10-24% of the general population and 57-74% of obese individuals are affected by nonalcoholic fatty liver disease (NAFLD). This can progress to nonalcoholic steatohepatitis

(NASH), a more life-threatening form of NAFLD in 3% of non-obese, 19% of obese and 50% of morbidly obese individuals (de Alwis and Day 2008). NASH is characterized by hepatocyte injury, inflammation and fibrosis which can lead to cirrhosis, liver failure and hepatocellular carcinoma. Because most patients with fatty liver do not get hepatitis, a two hit hypothesis was proposed for NASH with the first hit being hepatic steatosis and insulin resistance, and the second hit was oxidative stress coming from inflammation or exposure to a toxic xenobiotic.

The second hit was based on an increase in inflammation biomarkers in those patients with

26 hepatitis (Day and James 1998). Fructose metabolites, e.g. glyceraldehyde, seem to be involved as NASH was also associated with an approximate 40% sustained increase in serum advanced glycation end-product (AGE)-2 levels (Hyogo et al. 2007) formed from glyceraldehyde.

In the following, accelerated cytotoxic mechanism screening techniques (ACMS) was used in isolated rat hepatocytes with doses of xenobiotics sufficient to cause hepatocyte cytotoxicity in 2 hours. Previously this hepatocyte LC50 for 12 chlorobenzene analogues was shown to correlate with their in vivo hepatotoxicity in 24-36h (Chan et al. 2007). Although hepatocyte viability in vitro was not affected by 1 M fructose, hepatocyte cytotoxicity and mitochondrial toxicity occurred with 10-15mM fructose when the hepatocytes were exposed to continuous non-toxic oxidative stress as a model for liver inflammation (Lee et al. 2009).

Furthermore, 15μM glyoxal, a fructose Fenton oxidation product, also markedly increased hepatocyte susceptibility to H2O2 causing oxidative stress induced cell death (Shangari et al.

2006). In the following we have shown that fructose, and particularly glycolaldehyde (a fructose metabolite on the end-product oxalic acid forming pathway) was oxidized by hydroxyl radicals formed by the Fenton reaction (H2O2/Fe(II)) to form glyoxal which markedly increased protein carbonylation and increased hepatocyte cytotoxicity.

27 3.2. RESULTS

As shown in Table 1, there was at least a 70 fold increase in fructose cytotoxicity towards isolated hepatocytes when the hepatocytes were exposed to a non-toxic H2O2 generating system generated by glucose/glucose oxidase used as a cellular model for liver inflammation. Deferiprone (ferriprox), the ferric chelator used clinically to protect the liver from fructose/iron toxicity, prevented fructose/H2O2 cytotoxicity. Furthermore, loading the hepatocytes with 2µM Fe using ferrous iron and 8-hydroxyquionline complex (Fe(II):OHQ) markedly increased the H2O2 enhanced cytotoxicity, suggesting that the fructose was oxidized by a Fenton system (H2O2/Fe(II)) to a toxic product. Furthermore cytotoxicity was prevented by the hydroxyl radical scavengers, tryptophan and tyrosine.

28 Table 3.1. Fructose cytotoxicity is markedly enhanced by non-toxic H2O2/Fe(II) in isolated rat hepatocytes.

Treatment Cytotoxicity (% trypan blue uptake)

60 Min 120 Min 180 Min Control 17 ± 2 18 ± 3 21 ± 2 + Fructose 500mM 19 ± 2 18 ± 2 22 ± 2 + Fructose 15mM 21 ± 3 19 ± 3 23 ± 3 a a a,b + H2O2 38 ± 3 66 ± 7 92 ± 9 + Deferiprone 0.2mM 21 ± 3 b 39 ± 3 a,b 43 ± 4 a,b + Tyrosine 2mM 23 ± 3 b 37 ± 2 a,b 44 ± 2 a,b + Tryptophan 2mM 25 ± 2 b 42 ± 4 a,b 49 ± 3 a,b + Fe(II):OHQ 2µM 59 ± 6 a,b 92 ± 8 a,b 100 a,b + Fe(II):OHQ 2µM 25 ± 3 30 ± 4 a,b 31 ± 3 a,b + Fe(II):OHQ 2μM 20 ± 2 21 ± 3 23 ± 2

+ H2O2 22 ± 3 23 ± 2 24 ± 3 + Deferiprone 0.2mM 23 ± 2 24 ± 3 23 ± 3

Fe(II):OHQ, ferrous:8-hydroxyquinoline complex; H2O2 generating system, glucose 10mM and glucose oxidase 0.5 unit/ml. Mean ± S.E. for three separate experiments are given. a Significant as compared to control (P < 0.05). b Significant as compared to Fructose 15mM + H2O2 generating system (P < 0.05).

29 Table 2 shows that glycolaldehyde, a fructose metabolite, was cytotoxic with an LC50 after 2 hours exposure of about 20mM glycolaldehyde. Desferoxamine was also cytoprotective.

The LC50 2 hours decreased to 0.75mM glycolaldehyde when the hepatocytes were exposed to the non-toxic H2O2 generating system. Cytotoxicity was further increased by loading the hepatocytes with 2µM Fe(II):OHQ or cupric sulfate (Cu). Fe or Cu also increased glycolaldehyde cytotoxicity but the cytotoxicity was much less than when H2O2 was absent.

Glycolaldehyde by itself increased hepatocyte endogenous H2O2 levels as measured by the

FOX assay. The endogenous H2O2 was further increased when the H2O2 generating system is applied to glycolaldehyde, even though the H2O2 generating system alone did not increase hepatocyte H2O2 levels endogenously. Catalase inhibited hepatocytes by sodium azide were also 20 fold more susceptible than normal hepatocytes to glycolaldehyde.

30 Table 3.2. The cytotoxicity of glycolaldehyde, a fructose metabolite, is markedly enhanced by non-toxic H2O2/Fe(II) in isolated rat hepatocytes.

Cytotoxicity H2O2 Treatment 6 (% trypan blue uptake) nmol/10 cells

60 Min 120 Min 180 Min 90 Min Control 18 ± 2 20 ± 2 22 ± 2 5.2 ± 0.6 + Glycolaldehyde 20mM 30 ± 3 a 51 ± 4a 71 ± 6a 14.3 ± 1.3 a + Desferoxamine 0.2mM 26 ± 3 a,b 28 ± 3 a,b 35 ± 3 a,b 7.6 ± 0.7 a,b + Glycolaldehyde 1mM 21 ± 2 20 ± 2 22 ± 2 6.1 ± 0.6 a a a a + H2O2 42 ± 5 75 ± 7 100 34.0 ± 3.0 + Desferoxamine 0.2mM 24 ± 2 c 28 ± 3 a,c 35 ± 3 a,c 7.5 ± 1.0 c + Tryptophan 2mM 23 ± 3 c 31 ± 2 a,c 38 ± 3 a,c 8.0 ± 0.9 a,c + Aminoguanidine 1mM 29 ± 3 a,c 35 ± 3 a,c 39 ± 4 a,c 9.0 ± 1.2 a,c + Fe(II):OHQ 2µM 70 ± 6 a,c 100 a,c 100 39.0 ± 3.0 a + Cu 20µM 48 ± 5 a 83 ± 6 a 100 40.0 ± 4.0 a + Fe(II):OHQ 2µM 38 ± 3 a 58 ± 5 a 64 ± 6 a 8.2 ± 3.0 a + Cu 20µM 29 ± 3 a 32 ± 3 a 43 ± 5 a 8.1 ± 3.0 a a + H2O2 18 ± 2 22 ± 2 23 ± 2 7.2 ± 0.8 + Cu 20μM 19 ± 2 23 ± 2 24 ± 2 8.1 ± 0.7 a Catalase inhibited hepatocytes 22 ± 2 22 ± 3 23 ± 2 6.5 ± 0.5 + Glycolaldehyde 1mM 38 ± 4 a,d 52 ± 5 a,d 72 ± 7 a,d 21.0 ± 2.0 a,d

Fe(II):OHQ, ferrous:8-hydroxyquinoline complex, Cu, cupric sulfate; H2O2 generating system, glucose 10mM and glucose oxidase 0.5 U/ml; catalase inhibited hepatocytes, preincubating with sodium azide 4mM for 10min. Mean ± S.E. for three separate experiments are given. a Significant as compared to control (P < 0.05). b Significant as compared to Glycolaldehyde 20mM (P < 0.05). c Significant as compared to Glycolaldehyde 1mM + H2O2 generating system (P < 0.05). d Significant as compared to catalase inhibited hepatocytes (P < 0.05).

31 As shown in Table 3, a cell free system containing serum albumin was carbonylated by glyoxal about 15 fold more than by glycolaldehyde. The glycolaldehyde protein carbonylation rate was however increased several fold by Cu/H2O2 or Fe/ethylene-diamine-tetra-acetic acid

(EDTA)/H2O2. EDTA was used to complex iron and keep Fe(II) in a soluble state. A smaller increase was observed with glycolaldehyde and Cu or Fe/EDTA alone, whereas H2O2 alone did not increase glycolaldehyde protein carbonylation. Aminoguanidine prevented glycolaldehyde protein carbonylation induced by Fe(II)/EDTA/H2O2.

32 Table 3.3. Glycolaldehyde protein carbonylation is increased by Fenton’s reagent and inhibited by a glyoxal scavenger (aminoguanidine) in a cell free system.

Treatment Absorbance at 370nm 0 Min 180 Min Bovine serum albumin (BSA) 2mg/ml 0.11 ± 0.02 0.13 ± 0.01 + Glyoxal 0.1mM 0.50 ± 0.04 a 0.55 ± 0.05 a + Glycolaldehyde 0.5mM 0.15 ± 0.01 0.29 ± 0.03 a + Cu 20µM 0.18 ± 0.01 a,b 0.37 ± 0.04 a,b a + H2O2 1mM 0.15 ± 0.01 0.30 ± 0.04 a,b a,b + Cu 20µM + H2O2 1mM 0.18 ± 0.02 0.56 ± 0.06 a + Cu 20µM + H2O2 1mM 0.15 ± 0.01 0.16 ± 0.02 + Glycolaldehyde 1mM 0.14 ± 0.01 a 0.25 ± 0.03 a + Fe(II) / EDTA 0.5mM 0.20 ± 0.02 a,c 0.32 ± 0.06 a a a + H2O2 1mM 0.14 ± 0.01 0.19 ± 0.01 a,c a,c + Fe(II) / EDTA 0.5mM + H2O2 1mM 0.30 ± 0.03 0.92 ± 0.10 + Aminoguanidine 0.5mM 0.21 ± 0.02 a,c,d 0.53 ± 0.10 a,c,d a a + Fe(II) / EDTA 0.5mM + H2O2 1mM 0.28 ± 0.02 0.46 ± 0.05

Various agents were incubated with BSA (2mg/ml) in 50mM phosphate buffer (pH 7.4) at 37 °C. Protein carbonylation was determined by measuring DNPH-derivatized samples, measured at an absorbance of 370nm. Fe:EDTA, ferrous:ethylene-diamine-tetra-acetic acid complex; Cu, cupric sulfate; H2O2 generating system, glucose 10mM and glucose oxidase 0.5unit/ml. Mean ± S.E. for three separate experiments are given. a Significant as compared to control (P < 0.05). b Significant as compared to Glycolaldehyde 0.5mM (P < 0.05). c Significant as compared to Glycolaldehyde 1mM (P < 0.05). d Significant as compared to Glycolaldehyde 1mM + Fe/EDTA + H202 (P < 0.05).

33 3.3. DISCUSSION

Previously we showed that several drugs associated with idiosyncratic hepatotoxicity became much more toxic towards isolated hepatocytes if the hepatocytes were exposed to mild oxidative stress induced by a continuous low level reactive oxygen species generating system that did not affect cell viability or glutathione (GSH) levels. The toxicity of other drugs not associated with idiosyncratic hepatotoxicity were not affected by H2O2. This was used as a model to simulate isoniazid, hydralazine and amodiaquine induced hepatotoxicity mediated by activated neutrophils, macrophages and Kupffer cells, and was termed the hepatocyte inflammation model (Tafazoli et al. 2008; Tafazoli and O'Brien 2008; Tafazoli and O'Brien

2009). Previously, hepatic inflammation induced by bacterial endotoxin was shown to increase the in vivo hepatotoxicity of various drugs associated with idiosyncratic toxicity, e.g. chlorpromazine, diclofenac (Buchweitz et al. 2002; Deng et al. 2006; Lu and Cederbaum 2008).

This increased hepatotoxicity was attributed to activated immune cells that released cytokines, peroxynitrite, and reactive oxygen species generated by inducible nitric oxide synthase (iNOS) or NADPH oxidase (Morgan et al. 2008).

In the present study, the very high concentrations of fructose alone required for cytotoxicity is not physiological and probably resulted from excessive osmotic pressure resulting in cell dehydration. However, fructose induced hepatocyte toxicity was increased at least 70 fold if the hepatocytes were exposed to non-toxic levels of H2O2 generated by glucose/glucose oxidase. In rats receiving intraperitoneal fructose injection of 40µmol/g, plasma fructose level reached about 43 µmol/ml (~ 43mM) in 30 min (Morris, Jr. et al. 1978).

Therefore, the 15mM fructose required for 50% cytotoxicity in two hours in the hepatocyte

34 inflammation model could be considered within a physiological range. Cytotoxicity was also detectable if the hepatocytes were incubated for a longer time with 5mM fructose in the hepatocyte inflammation model (results not shown).

The cytotoxicity was also preceded by ROS and H2O2 formation that was prevented by desferoxamine, a ferric chelator. In addition, fructose/H2O2 cytotoxicity was further increased by adding trace amounts of ferric or cupric salts and suggested that hydroxyl radicals formed by

Fe/H2O2 (Fenton’s) reaction oxidized fructose or its metabolites to endogenous toxins.

Furthermore, cytotoxicity and ROS formation were prevented by the hydroxyl radical scavengers, mannitol, thiourea, and benzoate. When dehydroascorbate was used to permeate and load the hepatocyte with ascorbate (a reductant and antioxidant) the fructose/H2O2cytotoxicity and ROS formation was prevented (Lee et al. 2009). We have now shown that deferiprone, another clinically used ferric chelator, also prevented fructose/H2O2 cytotoxicity as did the hydroxyl radical scavenger tryptophan or tyrosine (Chetyrkin et al.,

2008). On the other hand Fe(II):OHQ used to permeate and load hepatocytes with Fe(II) markedly increased cytotoxicity. Previously, oxidized Fe(III):OHQ used to permeate and load hepatocytes with Fe(III) also markedly increased fructose/H2O2 cytotoxicity (Lee et al. 2009).

Fructose or glucose autoxidation is slow and requires an incubation time of weeks (Hunt et al. 1988). The mechanism is believed to involve a slow retroaldol condensation to an intermediary glycolaldehyde that is oxidized by superoxide radicals to form glyoxal

(Thornalley et al. 1999). Serum albumin showed some carbonylation with fructose that was 3-5 fold faster than glucose(Lawrence et al. 2008; Takagi et al. 1995). Fe or Cu increased fructose

35 autoxidation and was prevented by diethylenetriaminepentaacetic acid (DETAPAC), a transition metal chelator (Lawrence et al. 2008; Takagi et al. 1995). However fructose/H2O2 cytotoxicity was not inhibited by cell impermeable DETAPAC so cytotoxicity was not caused by extracellular fructose autoxidation (Lee et al. 2009).

Other researchers found that 1mM carbohydrates in phosphate buffer pH 7.4 were oxidized by 1mM H2O2 + 100µM Fe(II)/EDTA rapidly in 15 min to form glyoxal (using high performance liquid chromatography with fluorescence detection). The yields found were dihydroxyacetone (14%) > glyceraldehyde (12.6%) > glycerol(11%) > ribose (7.4%) > fructose

(5.8%) > glucose (4.5%) (Manini et al. 2006). Dihydroxyacetone 3mM at pH 8 in 30 min was also oxidized by these Fenton free radicals more readily than fructose at pH 4 to form glycolic acid (for both likely via a glycolaldehyde intermediate). Also formed was formate (from both), hydroxyl radicals (from dihydroxyacetone), and ribose (from fructose) (Maksimovic et al.

2006). Furthermore, 60% of 3mM glucose was oxidized by Fenton radicals at pH 5 in less than

3 hours to eight identified products including arabonic and gluconic acids (Buriova et al. 2004).

Fe also forms a complex with fructose (Lawrence et al. 2008) and the ferrous iron fructose complex could react with H2O2 to form a more powerful oxidizing agent that oxidizes fructose and especially its carbonyl metabolites.

Previously in our lab, it was showed that glyoxal toxicity towards hepatocytes occurred at 10µM in the hepatocyte inflammation model thereby representing a more than 200 fold increase in cytotoxicity (Shangari et al. 2006). Glyoxal could hence be the most likely endogenous toxin formed from fructose or its metabolites. Glyoxal is trapped by

36 aminoguanidine to form 3-amino-1,2,4-triazine (Thornalley 2003) which could explain why aminoguanidine prevented both fructose or glyoxal cytotoxicity towards the hepatocyte/H2O2 model.

The most genotoxic fructose metabolite formed from glycolysis is glycolaldehyde formed by glyceraldehyde oxidation catalyzed by cytosolic aldehyde dehydrogenase 1 (ALDH1) to form D-glycerate. This is then reduced to hydroxypyruvate (catalyzed by lactate dehydrogenase) which undergoes a decarboxylase catalyzed decarboxylation to release CO2 and form glycolaldehyde. Glycolaldehyde may also be formed from the oxidation of glyceraldehyde by the Fenton’s reaction. Glycolaldehyde is a mutagen, a protein DNA cross- linking agent, and the major metabolite of ethylene oxide, a class I human carcinogen.

Glycolaldehyde autoxidation is initiated by superoxide radicals to form glyoxal (Al-Maghrebi et al. 2003; Okado-Matsumoto and Fridowich 2000).

There was a 20 fold increase in glycolaldehyde cytotoxicity when the hepatocytes were also exposed to a non-toxic dose of H2O2. Furthermore this cytotoxicity was prevented by aminoguanidine (a glyoxal trap). Both glycolaldehyde/H2O2 and glycolaldehyde induced cytotoxicity were prevented by the Fe chelator desferoxamine Glycolaldehyde/H2O2 cytotoxicity was also markedly increased by Fe loading or by Cu, and was inhibited by tryptophan (a powerful hydroxyl radical scavenger) (Chetyrkin et al., 2008) indicating that cytotoxicity could be attributed to Fenton radicals. Glycolaldehyde also increased intracellular

H2O2 levels much more than the H2O2 formed by glucose/glucose oxidase which was presumably readily removed by intracellular GSH peroxidase and catalase. The glycolaldehyde

37 source of H2O2 is presumably the glycolaldehyde radical that readily reacts with oxygen to form glyoxal, hydroxyl radicals and H2O2 (Butkovskaya et al. 2006).

Glyoxal rapidly carbonylated serum albumin lysine, thiols and arginine residues (Glomb and Monnier 1995; Zeng and Davies 2005). We have shown that glycolaldehyde, carbonylated serum albumin more than 15 fold slower than glyoxal, but the glycolaldehyde carbonylation rate was markedly increased by Fe or Cu provided H2O2 was also present. This carbonylation was inhibited by aminoguanidine, suggesting that glycolaldehyde was readily oxidized by a

Fenton’s reaction to form glyoxal which bound to serum albumin. Other investigators however thought that glycolaldehyde was mostly oxidized by superoxide radicals to glyoxal (Al-

Maghrebi et al. 2003; Okado-Matsumoto and Fridowich 2000).

In Figure 3.1, the proposed biochemical mechanisms for the sequence of events that connect fatty liver (steatosis) to steatohepatitis are outlined based on the two hit hypothesis originally proposed by Day and James, 1998. The first hit would include the Western diet, characterized by refined sucrose, fructose or fat, perhaps in combination with insufficient levels of micronutrients. The second hit proposed would be inflammation, oxidative stress caused by hydrogen peroxide (H2O2) or ROS. ROS could be formed by the NADPH oxidase of the inflammatory cells; tumour necrosis factor alpha (TNF-α) induced mitochondrial toxicity, or drugs/xenobiotics P450 oxygen activation. Oxygen activation also occurs when oxygen reacts with glycolaldehyde radicals formed when fructose or fructose metabolites (glycolaldehyde) are oxidized by Fenton radicals (H2O2 + ferrous iron). The second hit can be prevented by glyoxal/methylglyoxal scavengers, lipid antioxidants, ROS scavengers or micronutrients.

38

Healthy Liver

zWestern diet: Refined SUCROSE, FRUCTOSE, fat zLow micronutrients zIncreased lipogenesis and free fatty acids 1st Hit zDecreased fatty acid β-oxidation zINSULIN RESISTANCE

Hepatic Steatosis (fatty liver)

zEnvironmental factors zDiet (FRUCTOSE) zXenobiotics or drugs zMitochondrial superoxide zOxidative stress (increased H2O2 levels)

zGlyoxal scavengers zROS formation (mitochondrial toxicity, P450, TNF- zLipid antioxidants 2nd Hit zROS/OH· scavengers α) zMicronutrients zIron catalyzed OH· formation, lipid peroxidation zFructose/carbonyl metabolites oxidized Fenton’s OH· to GLYOXAL

Steatohepatitis (NASH)

Figure 3.1. The proposed molecular hepatotoxic mechanisms associated with fructose induced NASH as per the two-hit hypothesis (modified from Lee et al., 2009)

39 CHAPTER 4

HEPATOCYTE STUDY OF THE MOLECULAR CYTOTOXIC MECHANISMS OF ENDOGENOUS TOXINS FORMED BY HYPEROXALURIA

4.1. INTRODUCTION

Hyperoxaluria is a condition characterized by increased endogenous oxalate and its increased excretion. Excessive oxalate crystallizes with calcium forming insoluble calcium oxalate crystals, and can accumulate to form kidney stones or block renal tubules. The elevated oxalate could arise from increased dietary oxalate intake (dietary hyperoxaluria) or increased absorption (enteric hyperoxaluria), or from elevated endogenous production of oxalate due to genetic defects in the detoxification of its immediate precursor glyoxylate (primary hyperoxaluria). Primary hyperoxaluria is an autosomal recessive disorder, and it is classified into two types. Primary hyperoxaluria type I (PH1) is due to a deficiency of peroxisomal alanine glyoxylate aminotransferase (AGT), which is responsible for glyoxylate transamination forming glycine, and requires pyridoxal phosphate as a cofactor. Deficiency of this enzyme would result in glyoxylate oxidation forming oxalate by glycolate oxidase or lactate dehydrogenase (LDH), and glyoxylate reduction forming glycolate by glycolate reductase (GR), also known as hydroxypyruvate reductase (HPR). Primary hyperoxaluria type II (PH2) is due to a deficiency of GR/HPR that catalyzes glyoxylate reduction to glycolate, and hydroxypyruvate reduction to D-glycerate. Deficiency of this enzyme leads to increased glyoxylate oxidation to oxalate catalyzed by glycolate oxidase or LDH, and hydroxypyruvate reduction to L-glycerate catalyzed by LDH.

40 Oxalate can be acquired directly from diet, food high in oxalate content includes chocolate, parsley, spinach, and rhubarb. Less than 15% of the daily oxalate intake is absorbed

(Voss et al. 2006). Absorption of unbound oxalate takes place along the entire gastrointestinal

(GI) tract including the colon through a passive non-energy requiring diffusion process (Binder

1974). Calcium and magnesium bind to oxalate, preventing it from being absorbed in the GI tract, and the product is excreted in the feces. Unabsorbed lipids, on the other hand, increase oxalate absorption by binding to calcium. The majority of urinary oxalate is synthesized endogenously in the liver as a metabolic end-product from sources such as dietary fructose and sucrose. Fructose was found to produce the greatest amount of oxalate compare to glucose and some sugar alcohols (Rofe et al. 1980). Therefore, chronic high consumptions of fructose and fat, such as seen in typical Western diets, may have contributed to the pathogenesis of . Furthermore, since the detoxification of glyoxylate to glycine by AGT is dependent on the cofactor pyridoxal phosphate, vitamin B6 deficiency could also lead to hyperoxaluria (Nishijima et al. 2003).

Oxalate has no known function in human and it cannot be further metabolized before it is excreted through the urinary system. It is filtered freely across the glomerulus (Asplin 2002), and is secreted at proximal tubule (Weinman et al. 1978). The normal excretion of oxalate in the is around 40mg daily (Taylor and Curhan 2008), whereas in the primary hyperoxaluria patients, it could be more than 200mg per day. Excessive endogenous oxalate production is problematic as oxalate readily crystallizes with calcium forming insoluble salts that precipitate in various tissues including the kidney, liver, heart, and joints (Friedman et al. 1962; Williams

1978). Accumulation of the crystals in renal system can cause stone formation, blockage of

41 renal tubules, inflammation and oxidative stress in renal epithelial cells (Scheid et al. 1996;

Thamilselvan et al. 2003). Treatments for hyperoxaluria include increasing urinary volume, high doses of vitamin B6, and supplements of orthophosphate or magnesium. Patients with primary hyperoxaluria develop renal failure at an early age, and a combined kidney and liver transplant is required eventually in order to treat the elevated endogenous production of oxalate.

Glyoxylate, a carboxylic acid and an aldehyde, is the immediate precursor of oxalate.

Glyoxylate can be formed through oxidation of its precursor, glycolate, by the enzyme glycolate oxidase which uses oxygen to produce hydrogen peroxide (H2O2). The precursor of glycolate, glycolaldehyde, can be obtained from metabolism of dietary fructose, or ethylene glycol oxidation through alcohol dehydrogenase. Other sources of glyoxylate include formation from glycine by D-amino acid oxidase, and from hydroxyproline, a major component in collagen. The detoxification of glyoxylate is through reduction to glycolate by cytosolic GR, or by transamination to glycine through AGT. In primary hyperoxalurias, reduced detoxification of glyoxylate will result in increased formation of its oxidation product, oxalate, by glycolate oxidase or LDH.

Hydroxypyruvate is known to be a precursor of oxalate (Rofe et al. 1980). It is formed from D-glycerate, the aldehyde dehydrogenase oxidation product of glyceraldehyde, a fructose metabolite. It readily undergoes decarboxylation to the genotoxic glycolaldehyde metabolite catalyzed by a decarboxylase (Hedrick and Sallach 1964). Glycolaldehyde is further oxidized to glycolate catalyzed by aldehyde dehydrogenase, then to oxalate via glyoxylate.

Hydroxypyruvate reduction to D-glycerate is dependent on NAD(P)H, catalyzed by the

42 ubiquitous cytosolic enzyme, hydroxypyruvate reductase (HPR). In PH2, deficiency in HPR leads to hydroxypyruvate reduction to L-glycerate through NAD(P)H dependent lactate dehydrogenase (LDH)

In calcium oxalate kidney stone disease, controversies exist over the endogenous toxins responsible for the toxicity. Some researchers have attributed the acute renal failure to glyoxylate induced toxicity and ATP depletion (Poldelski et al. 2001). Some other researchers however considered both oxalate and calcium oxalate monohydrate crystals were responsible

(Scheid et al. 1996; Thamilselvan et al. 2003). There is also a group of scientists who have attributed the toxicity solely to calcium oxalate monohydrate crystals, but not oxalate anion

(Guo and McMartin 2005; McMartin and Wallace 2005). The cytotoxic mechanism of primary hyperoxaluria is poorly understood, and requires a good model to study.

In the following, we have studied some metabolites of the fructose / oxalate pathway using primary rat hepatocytes. Metabolites being studied include: oxalate, the terminal product of the pathway; glyoxylate and hydroxypyruvate, whose metabolism is affected by the enzyme deficiencies in primary hyperoxaluria; and finally glycolate, the precursor for glyoxylate which accumulates in PH1.

43 4.2. RESULTS

Table 4.1 shows the doses of each of the fructose metabolites required to induce 50% of cell death at 2 hours (LC50 2 hours) measured by the trypan blue excusion assay in primary rat hepatocytes. Based on the LC50 2 hours doses, the toxicity rank was found to be glyoxylate > oxalate > hydroxypyruvate >> glycolate. However in the hepatocyte inflammation model where

H2O2 was produced by glucose and glucose oxidase system to mimic H2O2 generated by active immune cell NADPH oxidase (Tafazoli et al. 2008; Tafazoli and O’Brien 2009), the cytotoxicity fold increase found was glycolate (5.8 fold) > hydroxypyruvate (5.0 fold) > glyoxylate (2 fold) > oxalate (1.3 fold).

Table 4.1. Concentrations of each fructose metabolites studied that induced 50% cell death at 2 hours in isolated rat hepatocytes.

Approximate fold of increase Treatment LC 50 2 hours LC 50 2 hours with H2O2 (LC50 / LC50 with H2O2 ) Glyoxylate 8 ± 1 mM 4 ± 1.0 mM 2.0

Glycolate 23 ± 3 mM 4 ± 0.5 mM 5.8

Hydroxypyruvate 10 ± 2 mM 2 ± 0.5 mM 5.0

Oxalate 9 ± 2 mM 7 ± 1.0 mM 1.3

44 As shown in Figure 4.1, glyoxylate was toxic with a hepatocyte LC50 at 2 hours of 8 ±

1mM. Glyoxylate induced hepatocyte cell death was significantly increased by a non-toxic dose of Fe(III) > Cu(II), and was protected by Fe(III) chelator desferoxamine. Toxicity was also significantly increased by non-toxic H2O2 concentrations generated by glucose and glucose oxidase system (inflammation model), and by increasing endogenous H2O2 using sodium azide as a catalase inhibitor. Pyruvate, an endogenous H2O2 scavenger, protected against the cytotoxicity. Glyoxylate induced cytotoxicity involved its carbonyl group as it was protected by sulfur containing compounds, penicillamine and sulfite. Furthermore, glyoxylate toxicity was increased in the hepatocyte primary hyperoxaluria model when inhibitors of AGT or GR/HPR were used. Pyridoxal, one of the three vitaminers of vitamin B6, is a cofactor of many enzymes including AGT, it was shown to protect against cytotoxicity induced by glyoxylate. When LDH inhibitor was used, glyoxylate induced cytotoxicity was slightly increased.

45 120

a,b a,b a,b

) a,b 100 a,b a,b a 120min a a,b a,b 180min 80 a,b a,b a,b a,b a 60 a a,b b b b 40 b b b b b b

Cytotoxicity (% Trypan Blue Uptake 20

0 r r r M M M M r r µM ito ito ito ito trol ib m 5µ b n 8mM ) 4 1mM h 3 8mM ) 4µ 2 ib hi ibito e inhibitor l ) H2O2 h n h Co te (III II) 25µM + H2O2 in inhib in inhibitor la e ( e(III (II y se PR doxa F T x amin a DH DH in o + F x l Sulfite 10mM H ri L Cu AG L ly + Cu ta Pyruvate + R/ talase R/HPR i G + + AGT inhibitorG + + Py Ca G + Ca + Desfero + Penicillamine 4m +

Treatment

Figure 4.1. Glyoxylate induced cytotoxicity in primary rat hepatocytes measured by the trypan blue exclusion test at 120min and 180min. Fe(III):OHQ, ferric:8-hydroxyquinoline complex; Cu(II), cupric sulfate; H2O2 generating system, glucose 10mM and glucose oxidase 1 unit/ml; catalase inhibitor, sodium azide 4mM preincubated for 10 min; AGT inhibitor, 4mM propargylglycine preincubated for 30min; GR/HPR inhibitor, 10mM 2-carboxybenzaldehyde; LDH inhibitor, 40mM sodium oxamate. Mean ± S.E. for three separate experiments are given. a Significant as compared to control (P < 0.05). b Significant as compared to Glyoxylate 8mM (P < 0.05).

46 Table 4.2 shows the ROS formation and lipid peroxidation induced by glyoxylate.

Glyoxylate at its LC50 2 hours dose induced an increase in ROS formation at 60min and lipid peroxidation at 90min suggesting that oxidative stress was involved in the toxic mechanism.

With a non-toxic dose of Fe(III), glyoxylate induced ROS formation and lipid peroxidation increased significantly. Cu(II) also increased glyoxylate induced ROS formation and lipid peroxidation, but not as great when compared to Fe(III). Glyoxylate induced ROS formation increased in catalase inhibited hepatocytes, but not lipid peroxidation. H2O2 increased both glyoxylate ROS formation and lipid peroxidation. Both penicillamine and sulfite protected against glyoxylate ROS formation and lipid peroxidation. Furthermore, glyoxylate induced

ROS formation and lipid peroxidation was also increased by AGT inhibitor more than

GR/HPR inhibitor.

47 Table 4.2. Glyoxylate induced ROS formation in isolated rat hepatocytes.

ROS Lipid Peroxidation Treatment (F.I. units) (µM of MDA) at 60min at 90min Control 97 ± 4 0.40 ± 0.05 + Glyoxylate 8mM 215 ± 11a 0.98 ± 0.11 a + Fe(III):OHQ 4µM 368 ± 42 a,b 2.38 ± 0.25 a,b + Desferoxamine 1mM 110 ± 9 b 0.37 ± 0.04 b + Cu(II) 25µM 282 ± 21 a,b 1.93 ± 0.16 a,b a,b a,b + H2O2 267 ± 31 1.26 ± 0.14 + Catalase inhibitor 395 ± 35 a,b 1.19 ± 0.12 a + Pyruvate 3mM 142 ± 17 a,b 1.22 ± 0.14 a + Penicillamine 4mM 157 ± 13 a,b 0.47 ± 0.05 b + Sulfite 10mM 150 ± 16 a,b 0.54 ± 0.07 b + AGT inhibitor 412 ± 37 a,b 1.63 ± 0.14 a,b + GR/HPR inhibitor 276 ± 22 a,b 1.49 ± 0.13 a,b + Pyridoxal 8mM 108 ± 13 b 0.55 ± 0.05 b + LDH inhibitor 281 ± 30 a,b 1.37 ± 0.12 a,b + Fe(III):OHQ 4µM 95 ± 11 0.78 ± 0.09 + Cu(II) 25µM 73 ± 22 0.31 ± 0.03

+ H2O2 97 ± 8 0.37 ± 0.06 + Catalase inhibitor 88 ± 11 0.30 ± 0.06 + AGT inhibitor 86 ± 7 0.35 ± 0.05 + GR/HPR inhibitor 104 ± 11 0.37 ± 0.05 + LDH inhibitor 81 ± 12 0.39 ± 0.04

Fe(III):OHQ, ferric:8-hydroxyquinoline complex; Cu(II), cupric sulfate; H2O2 generating system, glucose 10mM and glucose oxidase 1unit/ml; catalase inhibitor, sodium azide 4mM preincubated for 10min; AGT inhibitor, 4mM propargylglycine preincubated for 30min; GR/HPR inhibitor, 10mM 2-carboxybenzaldehyde; LDH inhibitor, 40mM sodium oxamate. Mean ± S.E. for three separate experiments are given. a Significant as compared to control (P < 0.05). b Significant as compared to Glyoxylate 8mM (P < 0.05).

48 As shown in table 4.3, glyoxylate induced cytotoxicity, ROS formation, and lipid peroxidation was significantly increased by oxaloacetate. Glyoxylate undergoes a spontaneous condensation reaction with oxaloacetate forming oxalomalate, which is a competitive inhibitor of NADP+ dependent isocitrate dehydrogenase (Ingebretsen 1976) and aconitase in the citric acid cycle (Ruffo et al. 1959). When the hepatocytes were incubated with both glyoxylate and oxaloacetate, a decrease in mitochondrial membrane potential was observed possibly due to the formation of oxalomalate, which inhibited the citric acid cycle (Ingebretsen 1976) and the cellular antioxidant system (Ruffo et al. 1959), causing mitochondrial toxicity, ROS formation and lipid peroxidation. Glycine, a mitochondrial protective agent (Carini et al. 1997), was shown to protect hepatocytes against the cytotoxicity, ROS and lipid peroxidation.

49 Table 4.3. Endogenous glyoxylate reacts with mitochondrial oxaloacetate to form oxalomalate, a citric acid cycle inhibitor, which causes increase in ROS and lipid peroxidation, and collapse in mitochondrial membrane potential.

Cytotoxicity Mitochondrial Lipid Peroxidation ROS formation Treatment (% trypan blue uptake) membrane potential (µM of MDA) (F.I. units) at 120min (F.I. units) at 90min at 90min at 90 min Control 23 ± 2 487 ± 10 0.40 ± 0.03 98 ± 6 + Oxaloacetate 8mM 34 ± 3 a 501 ± 14 0.44 ± 0.05 119 ± 11 a + Glyoxylate 8mM 51 ± 4 a 520 ± 12 a 0.98 ± 0.11 a 283 ± 24 a + Oxaloacetate 8mM 70 ± 8 a,b 608 ± 17 a,b 1.95 ± 0.20 a,b 435 ± 33 a,b + Glycine 3mM 37 ± 4 a,b,c 542 ± 13 a,b,c 1.69 ± 0.14 a,b,c 162 ± 11 a,b,c + Glyoxylate 5mM 43 ± 4 a 511 ± 11 a 0.88 ± 0.09 a 188 ± 21 a + Oxaloacetate 5mM 57 ± 4 a,d 550 ± 11 a,d 1.91 ± 0.17 a,d 344 ± 31 a,d + Glycine 3mM 32 ± 3 a,d,e 515 ± 17 a,e 1.49 ± 0.13 a,d,e 202 ± 27 a,e

Mean ± S.E. for three separate experiments are given. a Significant as compared to control (P < 0.05). b Significant as compared to Glyoxylate 8mM (P < 0.05). c Significant as compared to Glyoxylate 8mM + Oxaloacetate 8mM d Significant as compared to Glyoxylate 5mM e Significant as compared to Glyoxylate 5mM + Oxaloacetate 5mM

50 Table 4.4 shows the LC50 2 hours dose of glycolate, the immediate precursor of glyoxylate, under various conditions. Glycolate was the least toxic among all the fructose metabolites studied with an LC50 2h of 23 ± 2mM. Its toxicity however was increased 46 fold with a catalase inhibitor (Siraki et al. 2002), and 6 fold in presence of H2O2 generated by glucose and glucose oxidase system (hepatocyte inflammation model). Glycolate toxicity was increased 2 fold in GSH depleted hepatocytes suggesting hepatic GSH peroxidase was involved in the detoxification of H2O2. Glycolate toxicity was also increased by Cu which likely catalyzed its autoxidation.

Table 4.4. Concentrations of glycolate that induce 50% cell death at 2 hours in isolated rat hepatocytes under various stress conditions.

Treatment LC50 2 hours concentration of glycolate

Glycolate 23 ± 2 mM + Cu(II) 25µM 12 ± 1 mM + GSH depleted 10 ± 1 mM

+ H2O2 4 ± 0.5mM + Catalase inhibited 0.5 ± 0.05mM

Mean ± S.E. for three separate experiments are given.

51 Figure 4.2 shows the hepatocyte cell death induced by hydroxypyruvate.

Hydroxypyruvate has an LC50 2 hours of 10 ± 2mM. Hydroxypyruvate induced cytotoxicity was significantly increased by Cu(II) > Fe(III). Transition metals likely catalyzed the autoxidation of hydroxypyruvate (Hedrick and Sallach 1961). Cytotoxicity was also increased by H2O2 and catalase inhibitor, and was partially protected by the H2O2 scavenger pyruvate.

Inhibitors of AGT and GR/HPR increased hydroxypyruvate induced cytotoxicity, but not as significantly as the increase in glyoxylate induced toxicity, possibly due to hydroxypyruvate being more upstream in the pathway. Pyridoxal protected against hydroxypyruvate induced hepatocyte cytotoxicity.

As shown in Table 4.5, hydroxypyruvate did not induce a significant increase in ROS formation at 2 hours. ROS formation was increased about 6 fold by non-toxic Cu(II), and 2.5 fold by non-toxic Fe(III). Cu also increased lipid peroxidation more than Fe. Since H2O2 can not directly oxidize DCFH-DA (Siraki et al. 2002; Zhu et al. 1994), FOX reagent was used and hydroxypyruvate was found to increase H2O2. H2O2 was further increased by Cu(II) > Fe(III), slightly increased in the inflammation model, and catalase and GR/HPR inhibited hepatocytes.

Desferoxamine and pyridoxal protected against the increase in H2O2 induced by hydroxypyruvate.

52 120

120min a,b a,b a,b a,b a,b a,b ) 100 a,b 180min a,b a 80 a a,b a,b a,b a,b a,b 60 a a,b a,b a,b b 40 b

Cytotoxicity (% Trypan Blue Uptake 20

0 r M 2 M to M M µM m µM O itor i tor trol m 5 2 b m ib bi n I) 4 H hi l 8m hi o 10 n te 3 inh n C ine 1 (II) 2 + a xa i Fe(II u e i o am C s ruv x la /HPR rid LDH ruvate + + ta Py py a + AGT inhibitor + y C + + Py ox + + GR ydr H + Desfero

Treatment

Figure 4.2. Hydroxypyruvate induced cytotoxicity in primary rat hepatocytes measured by the trypan blue exclusion test at 120min and 180min. Fe(III):OHQ, ferric:8-hydroxyquinoline complex; Cu(II), cupric sulfate; H2O2 generating system, glucose 10mM and glucose oxidase 1unit/ml; catalase inhibitor, sodium azide 4mM preincubated for 10 min; AGT inhibitor, 4mM propargylglycine preincubated for 30min; GR/HPR inhibitor, 10mM 2-carboxybenzaldehyde; LDH inhibitor, 40mM sodium oxamate. Mean ± S.E. for three separate experiments are given. a Significant as compared to control (P < 0.05). b Significant as compared to Hydroxypyruvate 10mM (P < 0.05).

53 Table 4.5. Hydroxypyruvate induced ROS formation and lipid peroxidation in primary rat hepatocytes.

H O formation ROS Lipid Peroxidation 2 2 (Absorbance Treatment (F.I. units) (µM of MDA) at 560nm) at 120min at 90min at 90min Control 97 ± 4 0.40 ± 0.03 0.089 ± 0.009 + Hydroxypyruvate 10mM 114 ± 12 0.95 ± 0.08 a 0.153 ± 0.012 a + Fe(III):OHQ 4µM 284 ± 21 a,b 1.16 ± 0.12 a,b 0.186 ± 0.020 a,b + Desferoxamine 1mM 103 ± 7 1.01 ± 0.09 a 0.103 ± 0.011 b + Cu(II) 25µM 664 ± 41 a,b 1.32 ± 0.14 a,b 0.245 ± 0.025 a,b a a a,b + H2O2 124 ± 10 0.97 ± 0.07 0.181 ± 0.014 + Catalase inhibitor 159 ± 17 a,b 1.04 ± 0.11 a 0.182 ± 0.015 a,b + AGT inhibitor 228 ± 26 a,b 0.98 ± 0.11 a 0.157 ± 0.016 a + GR/HPR inhibitor 112 ± 6 0.87 ± 0.13 a 0.190 ± 0.017 a,b + Pyridoxal 8mM 111 ± 8 1.08 ± 0.12 a 0.121 ± 0.015 a,b + LDH inhibitor 126 ± 7 a,b 1.03 ± 0.10 a 0.168 ± 0.014 a

Fe(III):OHQ, ferric:8-hydroxyquinoline complex; Cu(II), cupric sulfate; H2O2 generating system, glucose 10mM and glucose oxidase 1unit/ml; catalase inhibitor, sodium azide 4mM preincubated for 10min; AGT inhibitor, 4mM propargylglycine preincubated for 30min; GR/HPR inhibitor, 10mM 2-carboxybenzaldehyde; LDH inhibitor, 40mM sodium oxamate. Mean ± S.E. for three separate experiments are given. a Significant as compared to control (P < 0.05). b Significant as compared to Hydroxypyruvate 10mM (P < 0.05).

54 Figure 4.3 shows the cytotoxicity induced by oxalate, the terminal product in the pathway as it cannot be further metabolized. Oxalate though was toxic towards hepatocytes with an LC50 at 2 hours of 9 ± 2mM. Oxalate was slightly less toxic than glyoxylate and was more toxic than hydroxypyruvate. Toxicity was associated with increased ROS and lipid peroxidation formation (Table 4.6). Oxalate induced cytotoxicity, ROS and lipid peroxidation were increased by non-toxic doses of transition metal Cu(II)>Fe(III). Addition of an inhibitor of H2O2 detoxification by catalase increased oxalate cytotoxicity and oxidative stress, and

H2O2 generated by glucose and glucose oxidase system (hepatocyte inflammation model) increased oxalate cytotoxicity and ROS, but not lipid peroxidation. Equimolar calcium added to the hepatocytes with oxalate resulted in crystal formation observable under the microscope

(result not shown). Interestingly, those crystals did not seem to decrease cell viability.

Moreover, when calcium was added, ROS formation and lipid peroxidation were decreased compared to oxalate alone. When EDTA was added to chelate the calcium in the buffer (Guo and McMartin 2005), no significant change in oxalate induced cytotoxicity was observed, but

ROS formation and lipid peroxidation were decreased. We further tested calcium oxalate monohydrate crystals, and found it had a hepatocyte LC50 2 hours of 12mM, higher than that of oxalate. Calcium oxalate monohydrate induced a slight increase in ROS formation and lipid peroxidation, however, when compared to oxalate, it was significantly lower.

55 120

) a,b a,b 120min 100 a,b 180min a,b 80 a,b a,b a,b a a a,b a a a 60 a,b a a a a a a a a,b b a 40

20 Cytotoxicity (% Trypan BlueUptake

0 ) l M r ix * M o M o ly) tr m 4µM it e mM n 25µ b re m t 4 o H2O2 A C II) inhi (p ara T late 9 + e p u( s C mM (se Oxa + Fe(III) + 9 tala + BHA 50µM M + ED + Calcium 9mMm Ca iu + alc m 9m iu + C lc + Desferoxamine 0.5mM Ca +

Calcium oxalate monohydrate 12 m

Treatment

Figure 4.3. Oxalate induced cytotoxicity in primary rat hepatocytes measured by the trypan blue exclusion test at 120min and 180min. Fe(III):OHQ, ferric:8-hydroxyquinoline complex; Cu(II), cupric sulfate; H2O2 generating system, glucose 10mM and glucose oxidase 1unit/ml; catalase inhibitor, sodium azide 4mM preincubated for 10 min; BHA, butylated hydroxyanisole. Mean ± S.E. for three separate experiments are given. * Calcium was added 30 min after oxalate incubation. a P Significant as compared to control (P < 0.05). b P Significant as compared to Oxalate 9mM (P < 0.05).

56 Table 4.6. Oxalate induced ROS formation and lipid peroxidation in primary rat hepatocytes.

ROS Lipid Peroxidation Treatment (F.I. units) (µM of MDA) at 90min at 90min Control 97 ± 4 0.40 ± 0.03 + Oxalate 9mM 247 ± 23 a 1.81 ± 0.14 a + Fe(III):OHQ 4µM 535 ± 61 a,b 2.28 ± 0.21 a,b + Desferoxamine 0.5mM 56 ± 11 a,b 0.37 ± 0.04 b + Cu(II) 25µM 638 ± 58 a,b 3.72 ± 0.38 a,b a,b a + H2O2 353 ± 33 1.87 ± 0.19 + Catalase inhibitor 314 ± 29 a,b 2.29 ± 0.20 a,b + BHA 50µM 58 ± 7 a,b 0.35 ± 0.03 b + Calcium 9mM 163 ± 11 a,b 0.38 ± 0.04 b + Calcium 9mM (premix) 175 ± 22 a,b 0.35 ± 0.02 b + Calcium 9mM (separately)* 123 ± 15 a,b 0.30 ± 0.03 b + EDTA 98 ± 13 b 1.08 ± 0.15 a,b + Calcium oxalate monohydrate 12mM 122 ± 21 b 1.48 ± 0.17 a,b

Fe(III):OHQ, ferric:8-hydroxyquinoline complex; Cu(II), cupric sulfate; H2O2 generating system, glucose 10mM and glucose oxidase 1unit/ml; catalase inhibitor, sodium azide 4mM preincubated for 10 min; BHA, butylated hydroxyanisole. Mean ± S.E. for three separate experiments are given. * Calcium was added 30 min after oxalate incubation. a Significant as compared to control (P < 0.05). b Significant as compared to Oxalate 9mM (P < 0.05).

57 4.3 DISCUSSION

The endogenous toxins responsible for kidney stone disease have been attributed to a build-up of hepatic glyoxylate and oxalate. Much research has been carried out in the past studying primary hyperoxaluria, and there has been a debate as to which endogenous sugar metabolites were responsible for hyperoxaluria kidney toxicity. Free oxalate anion alone was found to increase free radical production in renal epithelial cells (Scheid et al. 1996). Some researchers have also found that both oxalate anion and calcium oxalate induced an increase in lipid peroxidation and ROS formation (Thamilselvan et al. 2000; Thamilselvan et al. 2003).

Another group of researchers have attributed the cytotoxicity induced by oxalate to the formation of calcium oxalate monohydrate crystals, and proposed that calcium oxalate alone was responsible for the cytotoxicity in renal epithelial cells (Guo and McMartin 2005). It was also proposed that calcium oxalate crystals, but not oxalate anion, increased superoxide radical formation (Khand et al. 2002), and induced mitochondrial permeability transition (McMartin and Wallace 2005). However other scientists found that when glyoxylate or glycolaldehyde were preincubated with isolated mouse proximal tubule segments, there was an induced ATP depletion and LDH release, but oxalate and glycolate did not induce toxicity. They have suggested that if calcium oxalate was toxic, toxicity would arise from crystal cast formation rather than a direct cellular cytotoxic effect (Poldelski et al. 2001). Since more than 85% of the urinary oxalate was suggested to be formed by endogenous metabolism (Baker et al. 1997), research with hepatocytes is critical in understanding the molecular cytotoxicity mechanism involved in primary hyperoxaluria.

58 In this study, the cytotoxicity of several fructose metabolites were examined, and based on the doses that caused 50% of cell death measured by trypan blue exclusion test in primary rat hepatocytes in 2 hours, the rank of toxicity from the most toxic to least toxic found was glyoxylate > oxalate > hydroxypyruvate >> glycolate. Inhibitors of AGT and GR/HPR were used to model the enzyme deficiencies in primary hyperoxaluria. Propargylglycine (PPG) was previously reported to be a good inhibitor of AGT, and 4mM PPG inhibited 90% of AGT activity in 20 min (Cornell et al. 1984). When PPG was used, glyoxylate and hydroxypyruvate induced cytotoxicity, ROS formation and lipid peroxidation were significantly increased. It was suggested that the majority of glyoxylate was detoxified by AGT to glycine (Williams

1978), and by inhibiting AGT, it was likely that this would lead to a build-up of glyoxylate.

This shift of metabolism would favour oxalate formation by LDH or glycolate oxidase. The

AGT inhibitor did not increase hydroxypyruvate toxicity as much as glyoxylate, possibly due to hydroxypyruvate being further upstream in the pathway. When 2-carboxybenzaldehyde

(CBA), an inhibitor of GR/HPR (Litvinovich and O'Brien 2007), was used, glyoxylate and hydroxypyruvate induced toxicity was increased, but not as much as when AGT inhibitor was used. This suggested that AGT was perhaps a more significant detoxification enzyme for glyoxylate. Pyridoxal phosphate is a cofactor for AGT and is essential in the detoxification of glyoxylate. In vivo rat studies showed that vitamin B6 deficient diet can lead to severe hyperoxaluria (Nishijima et al. 2006). Similar cases have also been reported in humans.

Vitamin B6 supplement has been used as a treatment option for hyperoxaluria and was effective in 10-30% of the patients (Nishijima et al. 2006). Pyridoxamine, one of the vitamers of B6 vitamin, has been shown to decrease crystal formation (Chetyrkin et al. 2005). In this

59 study, pyridoxal was also able to protect glyoxylate induced cytotoxicity and oxidative stress possibly by activating AGT and assisting glyoxylate reduction to glycine.

It was suggested that more than half of the urinary oxalate arises from glyoxylate metabolism (Williams 1978). In the present work, glyoxylate (8mM) induced hepatocytes cell death was preceded by ROS formation and lipid peroxidation thereby suggesting that oxidative stress contributed to the cytotoxicity. Non-toxic doses of transition metals Fe(III) and Cu(II) increased cytotoxicity, ROS formation, and lipid peroxidation induced by glyoxylate. The

Fe(III) chelator desferoxamine prevented glyoxylate toxicity, suggesting that transition metals possibly formed a pro-oxidant complex with glyoxylate. Glyoxylate is highly electrophilic due to its carbonyl group. Glyoxylate cytotoxicity can also be protected by nucleophilic sulphur containing compounds such as cysteine (Baker et al. 1994), penicillamine (Bringmann et al.

1992), and sulfite (Sharma and Schwille 1993) through adduct formation. L-cysteine, for example, forms thiazolidine-2,4-dicarboxylic acid with glyoxylate (Figure 4.4) (Baker et al.

1994). The adduct was not toxic, and was rapidly excreted when administered orally to rats

(Alary et al. 1989). In this study, both penicillamine and sulfite protected glyoxylate induced toxicity, ROS formation, and lipid peroxidation. Previously, some researchers suggested that penicillamine can protect against glyoxylate toxicity and oxalate production by adduct formation with glyoxylate (Bringmann et al. 1992), whereas others suggested that it formed complexes with pyridoxal phosphate, a cofactor for AGT, leading to decreased glyoxylate reduction to glycine and increased oxidation to oxalate causing toxicity (Baker et al. 1997). In patients with Wilson’s disease receiving penicillamine treatment, increased urinary oxalate was

60 observed, further suggesting that penicillamine might not be a suitable treatment option for hyperoxaluria (Hoppe et al. 1993).

O O O O OH S + HS OH HO OH N NH O 3 H Glyoxylate Cysteine Thiazolidine-2,4- dicarboxylic acid

Figure 4.4. Cysteine reacts with glyoxylate forming an adduct, thiazolidine-2,4-dicarboxylic acid, and decreases oxalate formation from glyoxylate (Baker et al. 1994).

Glyoxylate induced toxicity could also be attributed to the formation of oxalomalate, the non-enzymatic condensation product with oxaloacetate under physiological conditions (Figure

4.5) (Ruffo et al. 1959). Oxalomalate had been shown previously to induce an increase in intracellular ROS, lipid peroxidation protein and DNA modification (Yang and Park 2003). In this study, when glyoxylate and oxaloacetate were added to the hepatocytes, cytotoxicity, ROS formation and lipid peroxidation were significantly increased. A collapse in mitochondrial membrane potential was observed indicating mitochondrial damage. Oxalomalate competitively inhibited mitochondrial aconitase thus suppressing the citric acid cycle (Ruffo et al. 1959), and causing mitochondrial toxicity. It could also competitively inhibit NADP+ dependent isocitrate dehydrogenase (Ingebretsen 1976), an important source for endogenous

NADPH regeneration, thus compromising cellular defence against oxidative stress. Glycine protected against glyoxylate and oxaloacetate induced mitochondrial toxicity and oxidative stress possibly by preventing cytosolic Na+ overload and the increase of endogenous Cl-

(Carini et al. 1997).

61 O OH O O OH O OH + HO O O O OH OH O OH O Glyoxylate Oxaloacetate Oxalomalate

Figure 4.5. Glyoxylate condenses with oxaloacetate forming oxalomalate, an inhibitor of aconitase and NADP+ dependent isocitrate dehydrogenase.

Glyoxylate is oxidized to oxalate by LDH and glycolate oxidase. Glycolate oxidase also catalyzes glycolate oxidation forming glyoxylate, and the Km of LDH for glycolate is more than 15 fold lower than glyoxylate (Murray et al. 2008), suggesting that LDH is the major enzyme responsible for formation of oxalate from glyoxylate. When the LDH inhibitor sodium oxamate (Thornburg et al. 2008) was used, it was found that glyoxylate toxicity, ROS formation and lipid peroxidation were slightly increased, suggesting glyoxylate was more toxic than its oxidation product, oxalate.

Glycolate is the immediate precursor of glyoxylate. It is believed to have low toxicity.

Previous studies done using cultured Chinese hamster ovary cells which do not normally express AGT, GR/HPR, or glycolate oxidase, glycolate was found to be toxic only when glycolate oxidase was expressed, and toxicity was reduced when AGT or GR/HPR were expressed (Behnam et al. 2006), suggesting that glyoxylate was the toxic product of glycolate.

In isolated mouse proximal tubular segments, glycolate did not induce toxicity whereas the same concentration of glyoxylate induced ATP depletion and LDH release (Poldelski et al.

2001). Glycolate is oxidized to glyoxylate catalyzed by glycolate oxidase which uses oxygen and produces H2O2. When the catalase inhibitor, sodium azide (Silva and O'Brien 1989), was

62 used, the toxicity of glycolate and H2O2 formation was significantly increased suggesting that glycolate induced toxicity was due to its oxidation to glyoxylate, and catalase was crucial in the detoxification of the H2O2 formed. The cytotoxicity of glycolate was also markedly increased in GSH depleted hepatocytes, suggesting GSH peroxidase was also important in detoxifying H2O2 formed by glycolate oxidase.

Hydroxypyruvate is an important substrate, it is known to be a precursor of oxalate through decarboxylation to glycolaldehyde (Rofe et al. 1980) . In PH2, when there’s a deficiency of GR/HPR, hydroxypyruvate is reduced to L-glycerate by LDH instead of forming D-glycerate by HPR, leading to L-glycerate acidosis. Other researchers found that hydroxypyruvate was readily autoxidized in the presence of transition metals, with

Cu showing the greatest effect (Hedrick and Sallach 1961). We have found that hydroxypyruvate toxicity was markedly increased by Cu>Fe. Hydroxypyruvate did not induce a significant increase in ROS formation, except when there was non-toxic concentrations of

Cu(II)>Fe(III). Hydroxypyruvate induced an increase in H2O2 formation suggesting it was converted to glycolate via decarboxylation to glycolaldehyde, and the glycolate formed was oxidized to glyoxylate catalyzed by glycolate oxidase which also forms H2O2.

Transition metals Fe(III) and Cu(II) significantly increased oxalate induced toxicity,

ROS formation, and lipid peroxidation, suggesting that transition metals possibly formed pro- oxidant complexes with oxalate (Park et al. 1997). Calcium readily crystallizes with oxalate forming calcium oxalate. When administered to cells, calcium oxalate was taken up by an endocytosis process (Lieske et al. 1994) and accumulated inside the cell within 30 min of

63 administration (McMartin and Wallace 2005). Many researchers have found that calcium oxalate could induce cytotoxicity, free radical generation, and lipid peroxidation. In the present study, calcium was added to oxalate by different methods: directly, premixed with oxalate, or by preincubation. Calcium oxalate crystals were visible microscopically (result not shown), but did not increase oxalate induced toxicity, ROS formation or lipid peroxidation. We further studied calcium oxalate monohydrate, and found that it had a hepatocyte LC50 2 hours of

12mM, higher than that of oxalate alone when added to the hepatocytes. Calcium oxalate monohydrate increased ROS formation and lipid peroxidation, but not as much as when compared to oxalate alone. This suggested that formation of calcium oxalate was probably not the most significant cytotoxic mechanism for oxalate. Moreover, when we examined the toxicity of oxalate with EDTA, which could chelate calcium in the buffer (Guo and McMartin

2005), no significant difference in toxicity was observed. Oxalate induced ROS formation and lipid peroxidation was inhibited by EDTA, and this could be attributed to the metal chelating ability of EDTA. Oxalate induced cytotoxicity was likely due to complex formation with transition metals, rather than with calcium. Furthermore, calcium in the diet could be beneficial to reduce oxalate absorption, as less urinary and more fecal oxalate was observed with rats receiving a calcium enriched diet (Ribaya and Gershoff 1982). In a recent clinical trial, calcium enriched diet was shown to decrease urinary oxalate excretion and was proposed to protect against hyperoxaluria (Penniston and Nakada 2009) as it binds to oxalate in the gastrointestinal tract and prevents its absorption.

Figure 4.6 outlines a simplified pathway of oxalate formation from fructose. Toxicity of hydroxypyruvate, glycolate, glyoxylate, and oxalate towards primary rat hepatocytes were

64 compared in this study. We propose that both glyoxylate and oxalate are responsible for the cytotoxicity associated with primary hyperoxaluria. Glyoxylate induced toxicity could be attributed to its carbonyl group, or the formation of oxalomalate which suppresses the citric acid cycle. Oxalate induced toxicity could be attributed to oxalate forming pro-oxidant complexes with transition metals. Addition of calcium to oxalate did not increase oxalate induced cytotoxicity or oxidative stress unlike that reported previously by others.

65

Fructose

Abbreviations Fructose-1-phosphate ADH alcohol dehydrogenase AGT alanine glyoxylate aminotransferase Glyceraldehyde ALDH aldehyde dehydrogenase NAD+ GO Glyoxylate oxidase GR Glyoxylate reductase ALDH HPR Hydroxypyruvate reductase NADH HPDC Hydroxypyruvate decarboxylase D-Glycerate LDH Lactate dehydrogenase NAD(P)+ NAD+ DAO D-amino acid oxidase GR/HPR LDH NADHLDH NAD(P)H Hydroxypyruvate L-Glycerate NAD(P)H NAD(P)+ HPDC

CO2 ROS ADH Glyoxal Glycolaldehyde Ethylene glycol NAD+ NADH NAD+ ALDH NADH

NAD(P)+ Glycolate O2 GR/HPR GO NAD(P)H H O 4-Hydroxyproline Glyoxylate 2 2 Ascorbic Acid GO or LDH Pyridoxine NAD(P)+ AGT DAO NAD(P)H Calcium Oxalate Serine Glycine Calcium Oxalate (insoluble)

Figure 4.6. A simplified fructose to oxalate metabolic pathway (modified from (Danpure and Rumsby 2004)). PH1 results when the AGT gene is mutated, and PH2 results when the GR/HPR gene is mutated. Build-up of glyoxylate in primary hyperoxalurias would lead to increased oxalate formation

66 CHAPTER 5

CONCLUSIONS AND FUTURE PERSPECTIVES

5.1 GENERAL CONCLUSION

Our objective was to study the molecular cytotoxic mechanisms of fructose and some of its metabolites involved in NASH and primary hyperoxaluria. When fructose is absorbed, it is transported to the liver for metabolism. Due to the link between high chronic fructose intake and NASH hepatotoxicity, fructose toxicity in the liver was one of our primary goals of study.

The major end-point of fructose metabolism is oxalate, so high fructose or oxalic acid dietary intake can result in the formation of kidney stones. Primary hyperoxaluria occurs as a result of the inactivation of two hepatic enzymes that detoxify glyoxylate. This inactivation is caused by genetic mutations. Most research has focused on oxalate as an endogenous toxin which is mostly formed from fructose in the liver without causing liver toxicity but causes kidney toxicity. However, it is generally not realized that many of the intermediates on the glycolytic pathway from fructose to triose phosphates (the first part of the oxalate pathway) then produce endogenous carbonyl toxins, such as glyceraldehyde, methylglyoxal and glyoxal, that have been implicated in diabetes or Alzheimer’s disease. Other endogenous carbonyl toxins formed from glyceraldehyde include glycolaldehyde and glyoxylate before oxalate is formed. Therefore, we are also interested in examining the cytotoxicity mechanisms of these genotoxic and toxic carbonyl metabolites.

67 Molecular cytotoxic mechanism study of fructose and glycolaldehyde

Previously, we found that fructose was only toxic to rat hepatocytes at 1.5M, and the toxicity increased at least 70 fold in the inflammation model with non-toxic concentrations of

H2O2 produced by the glucose and glucose oxidase system (Lee et al. 2009). It was also found that toxicity was increased significantly by a non-toxic dose of Fe(III). In this study, we have found that Fe(II) also significantly increased fructose toxicity in the H2O2/hepatocyte inflammation model. The cytotoxicity was also inhibited by the clinically-used iron chelator deferiprone (ferriprox), as well as the hydroxyl radical scavengers tryptophan and tyrosine

(Chetyrkin et al. 2008). This suggested that fructose or its metabolites were oxidized by hydroxyl radicals formed by Fenton’s system (H2O2 and Fe(II)) to form a toxic product.

Next we studied the fructose metabolite glycolaldehyde as it is the most genotoxic fructose metabolite. It was found to have an LC50 2 hours dose of 20mM in rat hepatocytes.

However in the inflammation model, toxicity was increased more than 20 fold. Desferoxamine protected against glycolaldehyde toxicity in both untreated hepatocytes and in the inflammation model. Cytotoxicity was further increased by non-toxic concentrations of Fe and

Cu, which was inhibited by the iron chelator desferoxamine and the hydroxyl radical scavenger tryptophan (Chetyrkin et al. 2008), suggesting that toxicity could be attributed to hydroxyl radical formation by the Fenton’s system. Glycolaldehyde also caused an increase in endogenous H2O2 formation, and this effect was increased more in the inflammation model.

The control of H2O2 by itself did not increase endogenous H2O2 as it was likely rapidly detoxified by catalase, and catalase inhibited hepatocytes were 20 fold more susceptible to glycolaldehyde induced toxicity. Therefore, the source of glycolaldehyde H2O2 was

68 presumably the glycolaldehyde radical that readily reacted with oxygen forming glyoxal, hydroxyl radical, and H2O2 (Butkovskaya et al. 2006).

Glycolaldehyde was shown to autoxidize to glyoxal when initiated by superoxide radicals (Al-Maghrebi et al. 2003; Okado-Matsumoto and Fridovich 2000). Glyoxal was most likely the toxin formed from fructose as only 40µM was enough to cause significant hepatocyte death in the inflammation model (Shangari et al. 2006). In the cell free system, glyoxal carbonylated bovine serum albumin (BSA) 15 times faster than glycolaldehyde. However, glycolaldehyde protein carbonylation was increased several fold by Cu or Fe in the presence of

H2O2. Aminoguanidine, a well-known glyoxal trap (Thornalley 2003), protected against BSA protein carbonylation suggesting that glycolaldehyde was readily oxidized by Fenton’s reaction to form glyoxal which bound serum albumin.

In short, glyoxal was most likely the endogenous toxin responsible for fructose induced toxicity. Glyoxal formation from fructose was likely catalyzed by Fenton’s reaction.

Glycolaldehyde could form glyoxal through superoxide radical catalyzed autoxidation, or through the formation of glycolaldehyde radical by Fenton products of the H2O2 + Fe(II) reaction such as the hydroxyl radicals.

ACMS study of toxins involved in primary hyperoxaluria

The cytotoxicity associated with primary hyperoxaluria has been attributed to a build-up of glyoxylate and oxalate. Oxalate is an end-product so it is not further metabolized. However several researchers have suggested that oxalate and especially calcium oxalate monohydrate

69 crystals were toxic to renal epithelial cells and caused free radical formation and lipid peroxidation by an unknown mechanism ((McMartin and Wallace 2005; Scheid et al. 1996;

Thamilselvan et al. 2000). Another group of researchers found that oxalate was not toxic and suggested that toxicity in hyperoxaluria was due to glycolaldehyde and glyoxylate induced

ATP depletion (Poldelski et al. 2001).

In the present study, metabolites of fructose that could be involved in primary hyperoxaluria were added directly to the hepatocytes. One of the objectives was to rank the toxicity of those metabolites based on doses that cause 50% cell death in 2 hours in the hepatocytes. The toxicity rank found was glyoxylate > oxalate > hydroxypyruvate >> glycolate.

We also determined the toxicity of those fructose metabolites in the inflammation model.

Toxicity of all metabolites increased in the presence of non-toxic concentrations of H2O2 generated by glucose and glucose oxidase system, and toxicity fold increase found was glycolate (5.8 fold) > hydroxypyruvate (5.0 fold) > glyoxylate (2 fold) > oxalate (1.3 fold).

Next, we used enzyme inhibitors to model the enzyme deficiencies in primary hyperoxaluria and its related cytotoxicity. Propargylglycine (PPG), the inhibitors of AGT

(Cornell et al. 1984), and 2-carboxybenzaldehyde (CBA) the inhibitor of GR/HPR (Litvinovich and O'Brien 2007) were added to the hepatocytes together with glyoxylate or hydroxypyruvate.

Our results showed that glyoxylate cytotoxicity was significantly increased by the enzyme inhibitors, more than the increase in hydroxypyruvate toxicity, suggesting that a build-up of glyoxylate and its oxidation formation of oxalate caused the toxicity. Toxicity of both glyoxylate and hydroxypyruvate were increased more by AGT inhibitor than GR/HPR

70 inhibitor, suggesting that AGT was a more important detoxification enzyme compared to

GR/HPR, and the majority of glyoxylate was detoxified through reduction to glycine (Williams

1978).

Glycolate was shown to be the least toxic among all metabolites, the toxicity was increased about 46 fold in the presence of a catalase inhibitor and 2 fold in GSH depleted hepatocytes. This suggested that glycolate toxicity was probably due to its oxidation to glyoxylate by glycolate oxidase, which also forms H2O2 (Behnam et al. 2006). Catalase and

GSH peroxidase were found to be significant in detoxification of the H2O2 formed by glycolate oxidase.

Hydroxypyruvate had relatively lower toxicity in the hepatocytes compared to glyoxylate and oxalate. Its toxicity mechanism did not involve ROS formation, but an increase in H2O2 formation was observed suggesting hydroxypyruvate was presumably converted to glycolate via decarboxylation to glycolaldehyde, and the glycolate formed was oxidized to glyoxylate catalyzed by glycolate oxidase which also forms H2O2. Transition metals Cu and Fe likely catalyzed the autoxidation of hydroxypyruvate (Hedrick and Sallach 1961) causing oxidative stress and cytotoxicity.

The next stage of our objective was to evaluate the toxicity of glyoxylate and oxalate, as well as examine the effect of calcium on oxalate induced toxicity. Glyoxylate cytotoxicity was associated with oxidative stress, and toxicity was increased significantly by Fe>Cu. Glyoxylate induced toxicity involved its electrophilic carbonyl groups, as sulfur containing substances

71 penicillamine (Bringmann et al. 1992) and sulfite (Sharma and Schwille 1993) both protected against glyoxylate toxicity by adduct formation. Glyoxylate toxicity could also involve the formation of the condensation product of glyoxylate and oxaloacetate, oxalomalate, which causes mitochondrial toxicity and oxidative stress. Oxalomalate is a competitive inhibitor of aconitase in the citric acid cycle (Ruffo et al. 1959), and NADP+ dependent isocitrate dehydrogenase which is important for generating mitochondrial NADPH (Ingebretsen 1976).

Furthermore, oxalate was a little bit less toxic compared to glyoxylate, the toxicity was also associated with oxidative stress and was increased significantly by Cu>Fe. Transition metals likely formed pro-oxidant complexes with oxalate making it toxic (Park et al. 1997). Calcium addition did not increase oxalate induced toxicity or oxidative stress. Calcium oxalate monohydrate crystals were shown to have a higher LC50 2 hours compared to oxalate, further suggesting that calcium oxalate formation was probably not the major cytotoxic mechanism of oxalate induced cytotoxicity. Moreover, when an LDH inhibitor sodium oxamate (Thornburg et al. 2008) was used to block glyoxylate oxidation to oxalate, toxicity and ROS formation of glyoxylate were slightly increased suggesting glyoxylate was more toxic than oxalate.

In conclusion, toxicity in primary hyperoxaluria was most likely due to accumulation of both glyoxylate and oxalate, but not calcium oxalate. Glyoxylate induced toxicity likely involved its carbonyl group covalently reacting with protein thiol groups, as well as pro- oxidant complex formation with transition metals. Glyoxylate condensation with oxaloacetate also likely contributed to the cytotoxicity by interfering with the TCA cycle and cellular antioxidant system. Oxalate induced toxicity likely involved pro-oxidant complexes formation

72 with transition metals but not with calcium. Finally, calcium oxalate did not cause direct cellular cytotoxicity in the hepatocytes.

5.2 FUTURE PERSPECTIVE

NAFLD or NASH have been suggested to be the leading cause of liver dysfunction in

United States and Europe (Clark et al. 2003). Even though high fructose consumption has been linked with NASH (Collison et al. 2009; Ouyang et al. 2008), the cytotoxic mechanism of how high fructose contributes to the progression of NASH was still not identified in these papers.

As an in vitro model of NASH, at least the hepatocyte studies look at the actual site of damage, and freshly isolated primary rat hepatocytes have advantages over cultured hepatocytes as the enzyme activities are characteristic of those found in vivo. In the hepatocyte NASH model, we have combined high fructose and inflammation by using doses of fructose and H2O2 generated by glucose and glucose oxidase systems. However, the fact that the primary hepatocytes are only viable for a few hours implies this study might only reflect the acute events in NASH. The pathogenesis of NASH, however, is a long term event progressing from simple steatosis, to

NASH, then fibrosis and hepatocellualr carcinoma (McCullough 2004). Therefore, it is necessary to investigate the chronic toxicity of NASH and validate the results in vivo.

Previously, it was found that rats fed with a fructose enriched diet for five weeks developed hepatic fat deposits, an increase in hepatic triglyceride and cholesterol. This system was suggested to be a suitable model for human NASH study (Ackerman et al. 2005). Since

73 NASH is associated with inflammation, a source of oxidative stress is needed to stimulate inflammatory events involved in the progression from steatosis to NASH. An iron enriched diet could be used to induce oxidative stress (Lafay et al. 2005). Therefore, by feeding the rats with a high fructose and a high iron diet would likely induce symptoms and events resembling

NASH more efficiently. Once the in vivo model of NASH is better established, the toxicity induced by fructose can also be better determined. Markers of toxicity, in forms of oxidative stress and protein or DNA modification, as well as specific AGE levels can also be assessed.

The serum concentrations of fructose metabolites, glycolaldehyde and glyoxal, could be measured and analyzed. Finally, biomarkers of NASH in the in vivo rat model can be compared to those of NASH patients. If the mechanisms of fructose and inflammation induced

NASH is valid, this model could be used for the development and testing of potential therapeutic agents.

As for the hepatocyte model of hyperoxaluria, previous researchers have used cultured human hepatoma cell lines (Baker et al. 2004) to study the liver specific production of oxalate from glyoxylate. The present study in primary rat hepatocytes was beneficial in terms of variable control and effective modulation of the experimental conditions. Previous in vivo hyperoxaluria models have used ethylene glycol administration to induced renal stone formation (Thamilselvan and Menon 2005). Ethylene glycol is metabolized in the liver to form glycolaldehyde, followed by oxidation to glycolate, and finally to oxalate via glyoxylate. This model did not take into consideration the enzymatic deficiency in primary hyperoxaluria. There are identified inhibitors of glyoxylate detoxification enzymes that have been tested in vitro, but their effect in vivo is still unknown. Thus, propargylglycine (PPG) the inhibitor of AGT

74 (Cornell et al. 1984), and 2-carboxybenzaldehyde the inhibitor of GR/HPR (Litvinovich and

O'Brien 2007) could be tested in vivo. If they could be safely administered and induce appropriate enzyme inhibition, they would serve as a possible in vivo model of primary hyperoxaluria. Furthermore, glyoxylate injection can be used to induce hyperoxaluria instead of ethylene glycol (Okada et al. 2007). Once the in vivo model of hyperoxaluria is established, it can then be used for the development of potential therapeutics.

The most optimal treatment option for primary hyperoxaluria would be to find glyoxylate traps, thus preventing glyoxylate accumulation and oxalate production. The protective effect of N-acetylcysteine, sulfite, and other sulphur containing substances could be tested in vivo. Furthermore, 2-hydroxy-3-butynoic acid was shown to irreversibly inhibit glycolate oxidase purified from pea plants (Jewess et al. 1975). This compound might be beneficial in treating hyperoxaluria as it prevents glyoxylate formation from glycolate, as well as oxalate formation from glyoxylate.

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86 APPENDIX I: PERMISSIONS

87