REGULATION OF THE HUMAN XANTHINE OXIDOREDUCTASE GENE …ethesis.helsinki.fi/julkaisut/laa/kliin/vk/martelin/regulati.pdf · REGULATION OF THE HUMAN XANTHINE OXIDOREDUCTASE GENE Eeva

REGULATION OF THE HUMAN XANTHINEOXIDOREDUCTASE GENE

Eeva Martelin

Hospital for Children and AdolescentsUniversity of Helsinki

Finland

and

Program for Developmental and Reproductive BiologyBiomedicum HelsinkiUniversity of Helsinki

Finland

ACADEMIC DISSERTATION

Helsinki University Biomedical Dissertations No. 42

To be publicly discussed with the permission of the Medical Faculty of the University of Helsinki, in the Niilo Hallman Auditorium

of the Hospital for Children and Adolescents,on January 16th, 2004, at 12 noon.

Helsinki 2004

Supervisors

Professor Kari O RaivioHospital for Children and AdolescentsUniversity of HelsinkiHelsinki, Finland

Docent Risto LapattoHospital for Children and AdolescentsUniversity of HelsinkiHelsinki, Finland

Reviewers

Docent Pekka KallioOrion Corporation Orion PharmaTurku, FinlandandUniversity of HelsinkiHelsinki, Finland

Docent Ari RistimäkiUniversity of HelsinkiHelsinki, Finland

Opponent

Professor Bruce FreemanAnesthesiology, Biochemistry and Molecular GeneticsDirector, UAB Center for Free Radical BiologyUniversity of Alabama at BirminghamBirmingham, Alabama, USA

ISBN 952-10-1491-1 (printed version)ISBN 952-10-1492-X (PDF)ISSN 1457-8433

To my family

Table of Contents

4

TABLE OF CONTENTS

ORIGINAL PUBLICATIONS 7

ABBREVIATIONS 8

ABSTRACT 9

INTRODUCTION 11

REVIEW OF THE LITERATURE 13

Tissue and species-specific expression of xanthine oxidoreductase (XOR) 13

Human purine catabolism 13

Distribution of XOR in human tissues 15

Expression of XOR in other species 18

XOR: Structure and catalytic mechanism 19

Molybdoflavoenzymes 19

Active site of XOR 20

Iron-sulfur clusters and the FAD center 21

Xanthine dehydrogenase to oxidase conversion 21

Regulation of gene expression 23

Levels of regulation 23

Transcriptional regulators 26

Nuclear factor Y (NF-Y) 26

Gene regulation during development and in differentiated tissues 28

The XOR gene 30

Chromosomal localization of the XOR gene 30

Structure of the XOR gene 30

Regulatory regions of the XOR gene 31

Table of Contents

5

Regulation of XOR under different oxygen levels and by nitric oxide 33

Oxygen sensing and hypoxia-inducible factor 1 33

Other mechanisms of regulation of gene expression by oxygen 34

Regulation of XOR in hypoxia 36

Regulation of XOR in hyperoxia 38

Effects of nitric oxide on XOR activity 38

Regulation of gene expression by iron 39

Iron and ischemia-reperfusion injury 39

Post-transcriptional regulation by iron 40

Transcriptional regulation by iron 40

AIMS OF THE STUDY 42

MATERIALS AND METHODS 43

Isolation of XOR-specific clones 43

Somatic cell hybrid mapping panel 44

Polymerase chain reaction (PCR) 44

Fluorescence in situ hybridization (FISH) 44

Cell culture and exposures of cells 45

Transfections and reporter gene analysis 46

Assays of enzyme activities and iron measurement 47

Protein analysis 48

RNA analysis 49

Nuclear protein extracts 50

Electrophoretic mobility shift assay (EMSA) 51

Statistics 52

Table of Contents

6

RESULTS 53

Localization of the human gene for XOR on chromosome 2p22 53

Transcription factor NF-Y regulates the human XOR gene promoter 54

Transcriptional induction of XOR during enterocytic differentiation of Caco-2 colon carcinoma cells A role for NF-Y 55

Hypoxic induction of human XOR activity is mediated by a post-transcriptional mechanism 57

XOR expression is transcriptionally induced by iron, but XOR activity isdecreased by iron chelation at the post-translational level 59

DISCUSSION 61

Chromosomal localization of the human XOR gene 61

Regulation of the human XOR promoter 62

XOR expression during small-intestinal enterocyte-like cell differentiation 65

Regulation of XOR by oxygen and iron 66

Pathophysiological aspects 69

CONCLUSIONS 72

ACKNOWLEDGEMENTS 73

REFERENCES 75

Original Publications

7

ORIGINAL PUBLICATIONS

This thesis is based on the following original publications referred to in the text by their

Roman numerals:

I Rytkönen EMK, Halila R, Laan M, Saksela M, Kallioniemi O-P, Palotie A and Raivio

KO: The human gene for xanthine dehydrogenase (XDH) is localized on chromosome

band 2p22. Cytogenet Cell Genet 68:61-63, 1995

II Martelin E, Palvimo JJ, Lapatto R and Raivio KO: Nuclear factor Y activates the

human xanthine oxidoreductase gene promoter. FEBS Letters 480:84-88, 2000

III Martelin E, Lapatto R, Heikinheimo M and Raivio KO: Expression of xanthine

oxidoreductase is induced during enterocytic differentiation of Caco-2 cells. Submitted

IV Linder N, Martelin E, Lapatto R and Raivio KO: Post-translational inactivation of

human xanthine oxidoreductase by oxygen under standard cell culture conditions. Am

J Physiol Cell Physiol 285:C48-C55, 2003

V Martelin E, Lapatto R and Raivio KO: Regulation of xanthine oxidoreductase by

intracellular iron. Am J Physiol Cell Physiol 283:C1722-C1728, 2002

In addition, some unpublished data are presented.

Abbreviations

8

ABBREVIATIONS

AO aldehyde oxidase

AP-1 activator protein-1

ATF activating transcription factor

ATP adenosine triphosphate

ARNT aryl hydrocarbon receptor nuclear translocator

cAMP cyclic adenosine monophosphate

cDNA complementary DNA

cGMP cyclic guanosine monophosphate

C/EBP CCAAT enhancer binding protein

DCPIP 2,6-dichlorophenolindophenol

EMSA electrophoretic mobility shift assay

FAD flavin adenine dinucleotide

FISH fluorescence in situ hybridization

HIF hypoxia-inducible factor

HNF hepatocyte nuclear factor

IRE iron responsive element

I-R ischemia-reperfusion

IRP iron regulatory protein

mRNA messenger RNA

NAD+ nicotinamide adenine dinucleotide

NF- B nuclear factor B

NF-Y nuclear factor Y

ROS reactive oxygen species

RPA ribonuclease protection assay

RT-PCR reverse transcriptase-polymerase chain reaction

UTR untranslated region

VEGF vascular endothelial growth factor

XDH xanthine dehydrogenase

XO xanthine oxidase

XOR xanthine oxidoreductase

Abstract

9

ABSTRACT

Due to its ability to produce reactive oxygen species, xanthine oxidoreductase (XOR) has

been suggested to contribute to the pathogenesis of ischemia-reperfusion (I-R) injury.

Significant species differences in the expression of XOR exist. Therefore, an understanding of

its role in human pathology requires definiton of factors determining the expression of XOR

activity in human tissues under varying conditions.

The aim of this series of studies was to characterize the regulation of the XOR gene. We

started with the isolation of DNA clones corresponding to the human XOR gene and

proceeded to localize the gene on chromosome band 2p22. To study the transcriptional

regulation of the human XOR gene, DNA fragments corresponding to the XOR promoter

were cloned, reporter gene constructs created, and the function of the promoter studied in

transiently transfected cells. Furthermore, we attempted to identify transcription factors

involved in XOR regulation under normal cell culture conditions and during the

differentiation of an intestinal cell line. Mechanisms affecting the regulation of XOR by

oxygen and iron, both related to ischemia, were examined.

We obtained evidence for a functional role for the ubiquitous transcription factor nuclear

factor Y (NF-Y) in the regulation of the proximal XOR promoter. In colon carcinoma cells

undergoing small-intestinal enterocyte-like differentiation, an induction of NF-Y binding to

the XOR promoter was detected in parallel to the appearance of XOR activity, protein, and

messenger RNA. Transcription factors GATA-4 and GATA-6 were shown to bind to the

XOR promoter, but their functional role was not verified. Hypoxia did not activate XOR

transcription, but instead a post-translational modification of the active site of the XOR

protein was shown to account for the inactivation of XOR activity by high oxygen

concentrations and, conversely, for the induction of XOR activity during hypoxia. Increased

intracellular iron induced XOR expression at the transcriptional level, whereas iron chelation

reduced XOR activity by a post-translational mechanism.

In conclusion, our data indicate that the expression of the XOR gene in terms of enzyme

activity is regulated at multiple levels. Regulation of the human XOR promoter by NF-Y and

Abstract

10

the induction of XOR activity during enterocytic differentiation suggest an association of

XOR activity and cell differentiation. Our results on the mechanisms of XOR regulation by

oxygen do not conclusively establish the role of XOR in I-R injury, but provide an important

basis for future studies. The induction of XOR by iron may be another factor exacerbating I-R

injury.

Introduction

11

INTRODUCTION

The physiological role of xanthine oxidoreductase (XOR) is to catalyze the last two

oxidation reactions of purine degradation in man. In these reactions hypoxanthine is first

oxidized to xanthine and subsequently to urate. XOR exists in two forms, the dehydrogenase

(XDH) and the oxidase (XO) form, which use nicotinamide adenine dinucleotide (NAD+) and

O2 as the electron acceptors, respectively. Under physiological conditions, XDH is the

prevalent form of XOR. However, it has been hypothesized that in ischemia, concomitantly

with accelerated adenosine triphosphate (ATP) degradation and accumulation of substrates for

XOR, the XDH form converts into the XO form, which upon reperfusion produces superoxide

anion (O2 ) and hydrogen peroxide (H2O2) (Figure 1). These reactive oxygen species (ROS)

can further react to produce the extremely reactive hydroxyl radical (OH ) and peroxynitrite

(ONOO ). Despite their beneficial effects in host defense against micro-organisms and their

essential role in signal transduction, ROS are more frequently linked with adverse molecular

events, namely oxidative damage to protein, lipids, and DNA. Based on the ability of XOR to

produce ROS, it has been assigned a central role in the pathogenesis of ischemia-reperfusion

(I-R) injury. In addition, XOR has been associated, for example, with various heart and

vascular diseases and inflammation.

Even though XOR has been implicated in various pathological states, there is little direct

evidence of its importance to human pathology. The existing data are based mainly on the use

of XOR inhibitors, with putative unspecific effects, and on animal models. However, there are

major differences in the tissue distribution and activity levels of XOR between human and

other species. Therefore, conclusions based on the results obtained from animal studies

regarding the role of XOR in human pathology have to be made with caution. Furthermore,

only in man and higher primates is XOR the terminal enzyme of purine degradation, since all

other animals possess uricase to convert uric acid into allantoin.

In addition to the extent to which XOR produces ROS, factors determining the expression

of the human XOR gene and modulating XOR activity constitute the basis for the evaluation

of the significance of XOR in human pathology. The regulation of gene transcription,

messenger RNA (mRNA) stability, protein translation, and post-translational protein

Introduction

12

modifications may all contribute to the regulation of XOR activity. Little is known about the

regulation of the human XOR gene. In these studies, we have aimed at characterizing the

regulation of the XOR gene expression under varying conditions in order to better understand

its potential role in human pathology.

Figure 1. XOR in ischemia-reperfusion (I-R) injury. Substrates for XOR accumulate and the levels

of NAD+ decrease during tissue ischemia. It has been hypothesized that concomitantly the

dehydrogenase form (XDH) of XOR is converted into the oxidase form (XO), which produces reactive

oxygen species (ROS), O2 and H2O2, upon reperfusion. The in vivo significance of the potential of

XDH to use O2 as the electron acceptor and to produce ROS is unclear. The harmful effects of ROS

are mainly caused by oxidative damage to proteins, lipids, and DNA.

Review of the Literature

13

REVIEW OF THE LITERATURE

Tissue and species-specific expression of xanthine oxidoreductase (XOR)

Human purine catabolism

The purines, adenine and guanine, are essential components of nucleic acids, high energy

phosphates, and the key signaling molecules cyclic adenosine monophosphate (cAMP) and

cyclic guanosine monophosphate (cGMP). Furthermore, dietary ingestion contributes to body

pools of purines. De novo synthesis of purines from nonpurine precursors requires energy,

and therefore, under normal conditions, intermediates of purine degradation are efficiently

reutilized through purine salvage. However, in situations of increased ATP degradation and/or

decreased ATP synthesis, for example during tissue hypoxia or strenous exercise, in acutely

ill patients, and during cytolytic cancer treatment, the purine catabolic pathway is overloaded

by the accumulation of degradation products (Becker 2001). XOR catalyzes the last two

reactions of the purine catabolic pathway in man; the oxidation of hypoxanthine to xanthine

and the irreversible oxidation of xanthine to urate, to be excreted mainly by the kidney

(Figure 2).

In hereditary xanthinuria hypoxanthine and xanthine are not oxidized but accumulate in

body fluids, and the patients may present with symptoms of urinary tract stones or exercise-

associated muscle or joint pains. However, most of the patients are asymptomatic and their

prognosis seems to be unaffected by the defect in hypoxanthine and xanthine degradation. In

classical type I xanthinuria only XOR activity is lacking, whereas in type II xanthinuria both

XOR and aldehyde oxidase (AO), another molybdoflavoenzyme, activities are absent (Raivio

et al. 2001). In both types of xanthinuria the mode of inheritance is autosomal recessive

(Frayha et al. 1977). In patients with type I xanthinuria different mutations of the XOR gene,

either nonsense nucleotide substitution or deletion leading to early termination codon, have

been described (Ichida et al. 1997). On the other hand, in patients with type II xanthinuria, a

single base substitution in the gene coding for the molybdenum (Mo) cofactor sulfurase gene

has been identified (Ichida et al. 2001). This is in accordance with the deficiency of XOR as

well as of AO in these patiens, since both enzymes require the sulfur ligand to be attached to

the Mo for activity. In Mo cofactor deficiency, sulfite oxidase activity is missing in addition


14

to XOR and AO activities, and the clinical presentation of the patients is severe with

symptoms attributable to sulfite oxidase deficiency (Raivio et al. 2001).

Figure 2. Interconversions and degradation of purine nucleotides, nucleosides and purines. A

simplified overview of pathways leading to the formation of substrates for XOR is shown. The sources

of purines are depicted in circles. The reactions catalyzed by XOR and the reactions of purine salvage

pathway are illustrated. (Modified from Becker 2001 and Raivio et al. 2001.)


15

Distribution of XOR in human tissues

Critical for the hypothesis that XOR may have a role in human I-R injury is, whether XOR,

in its active form capable of generating ROS, is present in the human organ in question.

Therefore, several studies have attempted to determine the distribution of XOR enzyme

activity, protein, or mRNA in human tissues. The data are partially conflicting, possibly due

to the different methods of XOR determination (Parks and Granger 1986; Kooij 1994;

Harrison 2002). Data derived from studies using activity assay, immunohistochemistry or

mRNA assay to detect XOR in human samples are summarized in Table 1. Most of the

studies consistently indicate highest XOR expression, as assessed by enzyme activity, protein,

or mRNA levels, in the liver and proximal intestine (Vettenranta and Raivio 1990; Moriwaki

et al. 1993; Kooij 1994; Sarnesto et al. 1996; Saksela et al. 1998; Linder et al. 1999). Also the

mammary gland, especially during lactation, shows high expression of the XOR protein, and

XOR protein is found at high concentrations in human milk (Bruder et al. 1984; Sarnesto et

al. 1996; Linder et al. 1999). However, the specific activity of XOR in human milk is low

compared to the activity in the intestine and liver (Sarnesto et al. 1996), and both inactive

demolybdo- and desulfo-enzymes have been detected (Abadeh et al. 1992). On the other

hand, the high expression of XOR in the lactating mammary gland, and within the membrane

enveloping milk fat droplets, suggests an important structural role for XOR in milk secretion

(McManaman et al. 1999; McManaman et al. 2002; Vorbach et al. 2002). Interestingly, no

XOR protein was detected in malignant breast tumors (Cook et al. 1997).

Data concerning the lung and kidney are controversial, but suggest a markedly lower XOR

expression than in the liver and intestine. The presence of XOR in capillary and arteriolar

endothelium of several organs has been suggested (Bruder et al. 1984; Hellsten-Westing

1993; Moriwaki et al. 1993; Linder et al. 1999), and may explain the detection of XOR

mRNA by sensitive methods like reverse transcriptase-polymerase chain reaction (RT-PCR)

in organs with no visible immunoreactive XOR protein (Saksela et al. 1998). XOR has been

considered as a mainly cytosolic protein (Jarasch et al. 1981; Ichikawa et al. 1992), which is

in line with its physiological function in purine metabolism, but its localization also on the

surface of cultured endothelial cells has been reported (Rouquette et al. 1998; Harrison 2002).

While some studies have demonstrated XOR activity and antibodies in human serum (Bruder

et al. 1984; Spiekermann et al. 2003), others have failed to confirm these findings (Sarnesto et


16

al. 1996). However, it has been proposed that XOR may occasionally, for example due to

tissue damage in organs with high XOR content, be released into the circulation and bind to

endothelium, thereby contributing to distant organ damage (Pesonen et al. 1998; Houston et

al. 1999; Weinbroum et al. 1999).

Two studies have addressed the question of XOR activity and mRNA expression during

gestation (Vettenranta and Raivio 1990; Saksela et al. 1998). XOR activity is found in the

liver and intestine throughout gestation, and appears to increase in the liver and decrease in

the intestine toward term. Fetal brain, myocardium or kidney did not contain measurable

XOR activity, but low enzyme activity was detected in the lung. By using ribonuclease

protection assay (RPA), XOR mRNA was found in the developing liver and intestine, but not

in the myocardium, brain, lung or kidney. However, when the samples were analyzed by a

more sensitive method, RT-PCR, also brain, lung, and kidney revealed XOR mRNA (Saksela

et al. 1998).


17

Table 1. Expression of the XOR activity, protein, and mRNA in human tissues.

Reference XOR activity XOR protein XOR mRNA

Bruder et al.1984

milk lipid globulemembranes

IHCcapillary endothelial cells of heart, placenta, andkidney; sinusoidal cells of liver

ND

Hellsten-Westig1993

liver and skeletalmuscle extracts

IHC, ELISA, WBendothelial cells of capillaries and vascular smoothmuscle cells in cardiac and skeletal muscle;macrophages and mast cells in cardiac muscleneg: cardiac and skeletal muscle cells

ND

Moriwaki et al.1993

ND IHChepatocytes, sinusoidal cells of liver; enterocytes ofduodenum; arterial endothelium of heart, kidney,aorta, brain, lung and mesenteryneg: bile ducts

ND

Moriwaki et al.1996

ND IHCtongue, esophagus, stomach, trachea, salivary andsweat glands; mammary gland, liver, lung, heart,kidney, small and large intestine; uterus, prostate,striated muscle, pancreas, spleen, T-lymphocytesneg: aorta, mesentery, adrenal, ovary, urinary bladder,thyroid, B-lymphocytes, breast cancer metastasis

ND

Sarnesto et al.1996

liver, intestine,milk

WB, ELISAliver, intestine, milk, (lung, kidney) neg: brain, heart, skeletal muscle, serum

ND

Cook et al.1997

ND IHCmammary gland epithelial cellsneg: neoplastic mammary gland epithelium

ND

Saksela et al.1998

liver, intestine,lungneg: brain, heart,kidney

ND RPAliver, intestineRT-PCRbrain, lung, kidney,(heart)

Linder et al.1999

ND IHCliver; enterocytes and goblet cells of the proximalintestine; mammary gland epithelial cells; capillaryendothelium of mammary gland, skeletal muscle andkidney; endothelium of arterioles of mammary glandand kidneyneg: brain, heart, lung, skeletal muscle, kidney

ND

IHC, immunohistochemistry; ND, not determined; ELISA, enzyme-linked immunosorbent assay; WB, Westernblot analysis; neg, negative; RPA, ribonuclease protection assay; RT-PCR, reverse transcriptase-polymerasechain reaction


18

Expression of XOR in other species

In all mammals, XOR activity is highest in the liver and intestine. The measurable enzyme

activity and XOR protein in man is confined to relatively few organs, whereas most rat tissues

have detectable XOR activity (Hashimoto 1974; Parks and Granger 1986; Kooij 1994). In

mouse, the highest levels of XOR activity have been detected in the proximal small intestine,

lung, and liver and low levels in colon, kidney, spleen, and heart. The XOR protein was

present in the corresponding tissues, except for the lack of immunoreactive protein in the

kidney. XOR mRNA was strongly expressed in the small intestine, stomach, lung, and heart,

but at low levels in other tissues (Kurosaki et al. 1995). XOR has been purified from bovine

milk and localized to the epithelial and capillary endothelial cells of the mammary gland and

to capillary endothelium in many other bovine tissues (Jarasch et al. 1981).

In conclusion, compared to man, specific XOR activities, including serum activity, are

markedly higher and tissue distribution is wider in rodents and other mammals (Parks and

Granger 1986; Kooij 1994). Futhermore, there is a crucial difference in the purine catabolism

between man and higher apes, and other mammals. Namely, in other animals, uric acid is not

the end product of the purine catabolic pathway, but is further converted to allantoin by

uricase (Raivio et al. 2001). Consequently, the levels of urate, which acts as an antioxidant, in

the extracellular compartment are lower, which has been considered a disadvantage and a

possible reason for shorter life-span (Ames et al. 1981).

Mice with loss-of-function mutation in the XOR gene, have recently been generated. The

heterozygous XOR +/- mice appear healthy, but are unable to sustain lactation, which leads to

starvation and death of their offspring. XOR -/- mice die at an early age after birth, but their

phenotype has not been described in detail yet. However, based on the findings in the

heterozygous mice, XOR was assigned a structural role in milk fat droplet secretion (Vorbach

et al. 2002).


19

XOR: Structure and catalytic mechanism

Molybdoflavoenzymes

Molybdoflavoenzymes require both a Mo cofactor and flavin adenine dinucleotide (FAD)

for their catalytic activity. Only two human molybdoflavoenzymes, XOR and AO, have been

identified (Hille and Nishino 1995; Hille 2002; Garattini et al. 2003). The mammalian XORs

and AOs show high degree of homology at both gene and protein levels suggesting a common

evolutionary origin (Ichida et al. 1993; Terao et al. 1998; Terao et al. 2000). XOR catalyzes

the last two steps of the human purine degradation pathway, but the physiological function of

AO is unclear, even though endogenous substrates, including retinaldehyde and

dihydroxymandelaldehyde, have been identified. However, AO has been suggested to play an

important role in the metabolism of xenobiotics (Huang and Ichikawa 1994; Garattini et al.

2003). Whereas XOR occurs in two forms, the XDH form (EC 1.1.1.204), using preferably

NAD+, and the XO form (EC 1.1.3.22), using O2 as the electron acceptor (Hille and Nishino

1995), AO exists only in one form, which donates electrons to O2 (Turner et al. 1995). The

tissue distribution of the two human molybdoflavoenzymes differs especially with respect to

the intestine, where the expression of XOR is high (Sarnesto et al. 1996; Linder et al. 1999)

but that of AO low, and the lung, where the expression of AO is high (Wright et al. 1995), but

only very low XOR activity has been detected (Saksela et al. 1998).

Both XOR and AO are homodimers of identical 150-kDa subunits, which consist of a 20-

kDa N-terminal domain containing two iron-sulfur centers, a 40-kDa middle domain

containing FAD, and an 85-kDa C-terminal domain containing the molybdopterin cofactor

and the substrate binding site. The homodimeric enzyme has two identical active sites lined

by residues from both subunits. Divergence between the amino acid sequences of XOR and

AO around the FAD cofactor and the molybdopterin active site may account for the

differences in electron acceptor and substrate specificity of the enzymes, respectively (Calzi et

al. 1995; Enroth et al. 2000; Garattini et al. 2003).


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Active site of XOR

Based on crystallographic studies of bovine milk XOR (Enroth et al. 2000) and AO from

Desulfovibrio gigas (Romao et al. 1995; Huber et al. 1996), the structure and the function of

the active site of XOR have been elucidated. The crystal structures of bovine milk XDH

(Protein Data Bank (PDB) 1FO4) and XO (PDB 1FIQ) suggest that hypoxanthine and

xanthine bind to the C-terminal domain of XOR, close to the molybdopterin cofactor. During

substrate oxidation the Mo center of XOR is first reduced by electrons received from the

substrate and subsequently re-oxidized, as the electrons pass first to the iron-sulfur centers

and then to the FAD center, and are finally donated to NAD+ or O2 (Hille and Nishino 1995).

In the oxidized form of XOR, the ligands for the Mo ion are two pterin cofactor sulfurs, a

double-bonded sulfur atom, a double-bonded oxygen atom, and an oxygen atom with single

bond, whereas in the reduced form the double-bonded sulfur is substituted by a sulfhydryl

group (Figure 3) (Enroth et al. 2000). The oxygen atom that is attached to substrate during the

oxidation reaction is derived from a solvent water molecule. Recently, a mutation of the Mo

cofactor sulfurase gene has been detected as the cause for the absence of XOR and AO

activity in type II xanthinuria (Ichida et al. 2001). This sulfurase introduces a sulfur atom in

place of an oxygen atom at the Mo center of XOR and AO. Interestingly, an inactive desulfo

form of XOR, lacking the sulfur double bonded to the Mo, has been detected in rat liver cells

(Figure 3) (Ikegami and Nishino 1986). The sulfurase seems to have a physiological function

in regenerating the active sulfo form of XOR, which underlines the critical role of the sulfur

ligand at the Mo center. In addition, an inactive demolybdo form of XOR has been detected in

human milk (Abadeh et al. 1992).

Figure 3. Ligand binding to the Mo center of XOR. In the reduced form of XOR the double bonded

sulfur of the oxidized enzyme is replaced by a sulfhydryl group. In the desulfo-XOR the sulfur is

replaced by oxygen.

Mo

S

O

OHpterinsulfur

pterinsulfur

Mo O

O

OHpterinsulfur

pterinsulfur

Oxidized Reduced Desulfo-XOR

Mo

SH

O

Opterinsulfur

substrate

pterinsulfur


21

Iron-sulfur clusters and the FAD center

The N-terminal domain of XOR contains several cysteine residues critical for the binding

of the two non-identical iron-sulfur clusters. The Fe/S clusters are of the ferredoxin type

([2Fe-2S]) and differ from each other with respect to the electron paramagnetic resonance

signals and redox potentials (Enroth et al. 2000; Iwasaki et al. 2000; Nishino and Okamoto

2000). Transfer of electrons from the reduced Mo center to the Fe/S clusters is

thermodynamically favorable and is followed by electron flux to FAD.

The FAD cofactor is located in the middle domain of both XOR and AO, but differences in

the surrounding amino acid residues affect the properties of the two molybdoenzymes (Calzi

et al. 1995; Enroth et al. 2000). In the dehydrogenase form of XOR the electrostatic and

stereochemical environment of the FAD center favors the binding of NAD+ and consequently

final transfer of electrons to NAD+ instead of O2 under normal conditions (Harris and Massey

1997; Enroth et al. 2000, Kuwabara et al. 2003). However, under certain conditions XDH is

also able to use O2 as the electron acceptor, whereas due to conformational differences XO

cannot bind NAD+ at all (Enroth et al. 2000, Kuwabara et al. 2003).

Xanthine dehydrogenase to oxidase conversion

Under physiological conditions, XDH is the predominant form of XOR in tissues (Stirpe

and Della Corte 1969). During ischemia XDH is converted into XO, which upon

reoxygenation uses O2 as the electron acceptor. In the latter process, O2 and H2O2 are

formed. In ischemic tissues, ATP levels fall and the amount of XOR substrates increases.

Based on this, XOR has been assigned a role in the pathogenesis of ischemia-reperfusion

injury, the conversion of XDH to XO being a central part of this hypothesis (Figure 1) (Parks

et al. 1982; McCord 1985; Nishino 1994; Harrison 2002). However, although NAD+ is the

preferred electron acceptor under physiological conditions, XDH is also able to generate O2

and H2O2 by donating electrons to O2 when the levels of NAD+ are low (Hille and Nishino

1995; Harris and Massey 1997). The significance of ROS produced by XDH in vivo is not

clear, but favorable conditions prevail, for example, during reoxygenation following ischemia.

Studies performed with purified XOR (Stirpe and Della Corte 1969; Waud and

Rajagopalan 1976; Nishino 1994; McManaman and Bain 2002) and the crystal structures of


22

bovine milk XDH and XO (Enroth et al. 2000) suggest either a reversible XDH to XO

conversion by sulfhydryl oxidation or an irreversible conversion by proteolysis (Figure 4).

The exact mechanism and the extent to which the conversion occurs in mammalian tissues is

not clear. The thiol groups of conserved cysteine residues Cys535 and Cys992 are critical for

disulfide bond formation during conversion of XDH to XO, which can be reverted by

dithiothreitol and other reducing agents (Nishino 1997; Enroth et al. 2000; McManaman and

Bain 2002). However, it has been suggested that other as yet unidentified cysteine residues

crucial for the conversion exist (McManaman and Bain 2002). Proteolytic cleavage with

trypsin cleaves XDH after Lys-551 and with pancreatin after Leu219 and Lys569, located in

the linker segments between the subdomains of XOR, and results in irreversible conversion to

XO (Enroth et al. 2000). Despite cleavage of the polypeptide chains the fragments remain

united except under reducing conditions. The protease responsible for the XDH to XO

conversion in vivo has not been identified.

Figure 4. Xanthine dehydrogenase (XDH) to oxidase (XO) conversion. XDH can be converted into

XO either reversibly by sulfhydryl oxidation or irreversibly by proteolytic cleavage. The domains

corresponding to the iron-sulfur [2Fe-2S] center binding N-terminal, FAD binding middle, and Mo

cofactor binding C-terminal domain are depicted. In addition, the cysteine residues responsible for

disulfide bond formation during reversible conversion are shown. (Modified from Nishino 1994.)


23

Regulation of gene expression

Levels of regulation

The purpose of the regulation of gene expression is to assure temporally and spatially

accurate expression of proteins which enables cells, on one hand, to accommodate to the

changing requirements of the environment and, on the other hand, to undergo normal cell

growth and differentiation. Differences between organisms are mainly based on the evolution

of the regulatory networks that control gene expression, not on genes themselves, which are

often conserved between species (Hood and Galas 2003). To accomplish these goals, the

regulation of gene expression takes place at several levels (Orphanides and Reinberg 2002)

(Figure 5).

Chromatin is a complex of DNA and proteins that in dividing cells is packaged into

chromosomes. In non-dividing cells, chromatin is distributed diffusely throughout the nucleus

and appears as condensed heterochromatin or more open euchromatin. Chromatin structure is

related to gene expression and it is critically regulated by histones, the principal proteins of

chromatin, which can either promote or repress gene activation (Weintraub and Groudine

1976). Modifications of histones and their higher-order structures, nucleosomes, by chromatin

remodelling complexes determine whether a specific area of chromatin is active or inactive at

certain time point (Dillon and Festenstein 2002; Robertson 2002; Felsenfeld and Groudine

2003). Distinctive chromatin remodelling complexes either activate or suppress gene

transcription and may associate with coregulator proteins, which interact with proteins

binding directly to the regulatory elements of genes. Histone modifications include

acetylation, methylation, phosphorylation, and ubiquitination of amino acids. Reflecting the

complexity of the regulation of chromatin structure, several histone acetylases and

deacetylases have been identified (Neely and Workman 2002; de Ruijter et al. 2003).

Transcriptional regulation of gene expression depends on cis-acting DNA elements

interacting with trans-acting regulatory proteins, i.e. transcription factors. Enhancers,

silencers, and regulatory elements located in introns may further modulate transcription.

Ultimately, the transcriptional activation of a gene is determined by cross-talk between

chromatin remodeling enzymes and transcription factors.


24

Several mechanisms account for the regulation of gene expression at the mRNA level.

Capping, the addition of a GMP to the 5’-end of the transcript under synthesis and the

consequent methylation of the GMP, protects the nascent RNA from degradation, enables the

translocation of mRNA from the nucleus to the cytoplasm, and is involved in translation of

mRNA into protein. Splicing out of introns and the modification of the 3’-end of the pre-

mRNA further contribute to the processing of the translatable mRNA (Proudfoot et al. 2002).

The stability of mRNAs may change under varying conditions due to the binding of

regulatory proteins. Different mRNA sequence elements have been identified, which either

stabilize or destabilize the mRNAs carrying them, or bind proteins that affect translation

(Addess et al. 1997; Xu et al. 1997; Davis et al. 2001). In addition to regulatory proteins, a

growing number of non-coding RNAs (ncRNA) with regulatory functions on different levels

of gene expression have been discovered. The ncRNAs may stabilize mRNAs or target them

for degradation, regulate splicing, or have an effect on mRNA translation (Storz 2002).

Post-translational protein modifications, including carboxylation, hydroxylation,

acetylation, phosphorylation, methylation, cleavage, oxidation-reduction, and the addition of

cofactors, may contribute to the generation of active protein molecules. Furthermore, the

degradation of proteins, including transcription factors, is regulated, for example, by the

ubiquitin-proteasome system (Muratani and Tansey 2003). The inactive demolybdo and

desulfo forms of XOR are examples of enzyme forms lacking the essential cofactor or ligand

of the active center, respectively, and may represent another way of regulating the expression

of the XOR gene.


25

Figure 5. Regulation of gene expression at different levels. Chromatin remodeling through histone

modifications determines the structure and the activity of chromatin (1). Gene transcription is

regulated by transcription factors (TF) and coregulators recruiting RNA polymerase II to gene

promoters. ATG depicts the translational initiation site (2). The stability and translation of mRNAs is

regulated by RNA binding proteins and non-coding RNAs (ncRNA) constituting the post-

transcriptional level of gene regulation (3). At the post-translational level proteins are modified, for

example, by the addition of cofactors and by protein hydroxylation and phosphorylation, or may

undergo degradation (4).


26

Transcriptional regulators

The critical role of regulatory proteins, including transcription factors, controlling gene

expression has become evident along with the sequence analysis of the human genome. More

than 3000 of the 30 000 35 000 protein coding genes of our genome code for proteins

involved in transcription and translation (International Human Genome Sequencing

Consortium 2001).

As signals from the environment reach the nucleus, relevant transcription factors co-

operatively bind to the promoters of genes encoding proteins required for the appropriate

cellular response. Promoter-bound transcription factors then recruit the components of the

basal transcription machinery, the general initiation factors TFIIB, TFIID, TFIIE, TFIIF and

TFIIH, and RNA polymerase II, to the DNA. After the assembly of this preinitiation complex,

an ATP-dependent unwinding of the DNA, catalyzed by DNA helicase, takes place at the

transcriptional start site. Subsequently, the synthesis of RNA transcript commences (Dvir et

al. 2001). Individual sequence specific transcription factors interact with each other and the

basal transcription machinery either directly or through coregulator proteins (coactivators or

corepressors) to form multi-protein complexes, which mediate chromatin remodeling activity

or regulate the activity of the basal machinery (Martinez 2002). To terminate transcription,

transcriptional activators can be degraded, or their subcellular localization and interaction

properties can be altered by mechanisms involving ubiquitylation (Tansey 2001, Muratani and

Tansey 2003) and sumoylation (Seeler and Dejean 2003), respectively.

Nuclear factor Y (NF-Y)

Nuclear factor Y (NF-Y) is a ubiquitously expressed transcription factor binding to the

DNA sequence motif CCAAT. Several transcription factors can potentially bind to the

CCAAT-box, which is over-represented in eukaryotic promoter sequences and present in 30%

of them, but only NF-Y requires all five nucleotides (Bucher 1990; Mantovani 1998). Several

lines of evidence suggest a role for NF-Y in facilitating the recruitment of upstream DNA-

binding transactivators by stabilizing them and interacting with them (Wright et al. 1994), and

mediating interactions between upstream activators and the components of the basal

transcription machinery. NF-Y has been shown to interact with the TATA-binding protein

(TBP) and TBP-associated factors (TAFs), which mediate the interaction between upstream


27

activator elements and the basal transcription machinery by recognizing TATA or initiator

elements in the proximal promoters (Bellorini et al. 1997; Frontini et al. 2002). Furthermore,

the NF-Y binding site is often present in proximal promoter regions close to the sites which

bind the components of the basal transcription machinery.

NF-Y is a heterotrimer composed of three subunits; NF-YA, NF-YB, and NF-YC, the

latter two with similarity to histones. As a matter of fact, NF-Y is able to interact with

nucleosomes, thereby affecting the state of chromatin and promoting binding of other

transcriptional activators (Motta et al. 1999; Coustry et al. 2001; Romier et al. 2003). Histone

acetyltransferases GCN5 and P/CAF (Currie 1998a), and the chromosomal high mobility

group protein HMG-I(Y) (Currie 1997) have been shown to interact with NF-Y and thereby

possibly aid in the remodelling of chromatin structure. Upon binding NF-Y bends DNA,

which may further enhance transcriptional activation by allowing individual factors to come

to close vicinity with each other (Liberati et al. 1998).

The specificity of transcriptional responses involving ubiquitous transcription factors, like

NF-Y, is achieved through interaction with other factors that bind to the unique set of cis-

regulatory sequences of an individual promoter and are induced by a specific signal from the

environment or have cell-type restricted expression (Merika and Thanos 2001). Interactions

and co-operative binding of NF-Y with Sp1, Sp3 (Yamada et al. 2000), sterol regulatory

element binding protein (SREBP) (Dooley et al. 1998), hepatocyte nuclear factor-4 (HNF-4)

(Ueda et al. 1998) and CCAAT/enhancer binding protein (C/EBP) (Milos and Zaret 1992)

have been described. Furthermore, NF-Y modulates the fuction of the cAMP response

element binding protein (CREB) (Eggers et al. 1998) and interacts with the coactivator

proteins CBP (CREB-binding protein) and p300 (Faniello et al. 1999).

Besides being a ubiquitous transcription factor, regulating a variety of genes with different

functions and playing a crucial role as a promoter organizing factor, NF-Y participates in the

regulation of cell cycle progression by repressing the activity of cyclins, cyclin-associated

proteins, and a cyclin-dependent kinase (Hu and Maity 2000; Manni et al. 2001). In this

process NF-Y interacts with the tumor suppressor protein p53 (Yun et al. 1999). Moreover,

interaction of NF-Y with the proto-oncogene c-Myc has been described (Taira et al. 1999;


28

Izumi et al. 2001). NF-Y has been ascribed a role in hemoglobin synthesis (Liberati et al.

1998), and it is associated with myeloid (Marziali et al. 1997; Marziali et al. 1999; Sjin et al.

2002) and lymphocyte (Currie 1998b) as well as enterocyte differentiation (Bevilacqua et al.

2002).

Gene regulation during development and in differentiated tissues

Development of different cell types, organs, and ultimately different organisms is based on

the hereditary information carried in the genome. The key regulatory proteins in development

have been especially well conserved during evolution, and it is the genomic regulatory

network that mainly determines the unique characteristics of various organisms (Davidson et

al. 2002). The fundamental difference between developmental transcriptional responses

compared to physiological transcriptional responses is the progressivity of the former.

Cellular differentiation during development is influenced by maternal regulatory molecules,

inter- and intracellular signaling molecules, and the cis-regulatory sequences of the

responding genes. Changes in chromatin structure present another, epigenetic level of

regulation of gene expression during development and enable the differentiated, or

determined, cells to preserve their distinct gene expression patterns (Cunliffe 2003).

Among factors accounting for tissue-specific gene regulation are tissue-restricted

transcription factors. With respect to XOR, factors determining gene expression in the liver,

intestine, and mammary gland, which show the highest expression of XOR in human, are of

special interest. Transcription factors involved in liver-specific gene expression include the

families of homeodomain containing HNF-1, winged helix HNF-3, nuclear orphan receptor

HNF-4, and leucine zipper C/EBP proteins (Mendel and Crabtree 1991; Cereghini 1996;

Lekstrom-Himes and Xanthopoulos 1998; Zaret 2002). The expression of HNFs, which were

originally identified as factors regulating liver gene expression, is not restricted to liver, but

they contribute, for example, to the development of the gut (Shivdasani 2002). HNF-1

responsive element is essential for the transcriptional regulation of the small-intestinal

enterocyte-specific sucrase-isomaltase gene, and the ratio of HNF-1 to HNF-1 may affect

the expression of sucrase-isomaltase during intestinal development (Boudreau et al. 2001).

HNF-4 is a key regulator of hepatocyte differentiation, and among its target genes is HNF-1

(Zaret 2002). HNF-4 has also been assigned a crucial role in the regulation of apolipoproteins


29

AI and CIII in the intestine (Fraser et al. 1997). Interestingly, HNF-4 has been shown to

directly associate with NF-Y to enhance transcriptional activation (Ueda et al. 1998). Another

transcription factor, HNF-6, has been shown to regulate the differentiation of hepatoblasts

into biliary epithelial cells (Clotman et al. 2002).

The C/EBP transcription factor family consists of at least six members with different

patterns of tissue expression and functional consequences. C/EBP proteins can potentially

bind to the same CCAAT consensus sequence as NF-Y (Mantovani 1998) and are considered

critical for normal cellular differentiation and function (Lekstrom-Himes and Xanthopoulos

1998). C/EBP , - , and - are expressed in the liver and intestine among few other tissues.

During hepatocyte differentiation, the expression of new genes is often under the control of

C/EBP (Zaret 2002). Studies on the promoter of serum albumin, abundantly expressed in the

liver, have indicated transcriptional synergism between precisely positioned C/EBP and NF-Y

(Milos and Zaret 1992). C/EBP is induced by lipopolysaccharide, IL-6, IL-1, dexamethasone

and glucagon, suggesting a role in inflammatory response (Alam et al. 1992; Lekstrom-Himes

and Xanthopoulos 1998), which is also accompanied by up-regulation of XOR expression

(Dupont et al. 1992; Falciani et al. 1992; Pfeffer et al. 1994; Kurosaki et al. 1995; Chinnaiyan

et al. 2001). Consistent with a regulatory role in cellular differentiation and function, the

expression of C/EBPs in the mammary gland varies during pregnancy, lactation and

involution (Doppler et al. 1995; Gigliotti and DeWille 1998; Sabatakos et al. 1998).

The six members of the GATA transcription factor family share a highly conserved DNA

binding domain, which contains two zinc fingers and recognizes an (A/T)GATA(A/G) cis-

acting element in the promoters of various genes (Molkentin 2000; Patient and McGhee

2002). GATA-1, -2, and -3 are principally expressed in hematopoietic cells (Orkin 1992),

whereas GATA-4, -5, and -6 are expressed in most tissues of endodermal or mesodermal

origin and participate in development- and tissue-specific transcriptional gene regulation

(Arceci et al. 1993; Laverriere et al. 1994; Ketola et al. 2000; Fujikura et al. 2002). GATA-4,

-5, and -6 are all expressed in the mouse intestine, and GATA-4 has also been ascribed a role

in liver-specific gene regulation (Bossard and Zaret 1998). GATA-4 is considered a critical

regulator of the development of the gut and tissues derived from gut endoderm (Zaret 1999;

Shivdasani 2002). Interestingly, HNF-3 and GATA-4 are able to bind to their DNA binding


30

sites in compacted (inactive) chromatin in liver precursor cells, thereby initiating the events

leading to the opening of chromatin and facilitating the binding of other transactivators

(Cirillo et al. 2002). Functional co-operativity between GATA-5 and HNF-1 has been

proposed to mediate activation of intestinal gene promoters (Krasinski et al. 2001; van

Wering et al. 2002). Furthermore, GATA-4 and HNF-1 , together with the Cdx2

homeodomain transcription factor, regulate the intestine-specific sucrase-isomaltase gene

(Boudreau et al. 2002).

The XOR gene

Chromosomal localization of the XOR gene

The human gene for XOR was initially assigned to chromosome 2 by using a

complementary DNA (cDNA) clone as a probe in spot blot hybridization experiments (Ichida

et al. 1993). Subsequently, three research groups localized the XOR gene to chromosome

2p22 or 2p23, utilizing human-hamster hybrid cell lines and fluorescence in situ hybridization

(FISH) with human cDNA or genomic clones as probes (Xu et al. 1994b; Minoshima et al.

1995; Rytkönen et al. 1995). As the sequence of the human genome was published, the

location of the human XOR gene was confirmed on chromosome band 2p23.1 (International

Human Genome Sequencing Consortium 2001).

Consistent with the location of the human XOR gene, the chromosomal position of the

mouse gene for XOR has been mapped to distal mouse chromosome 17 containing gene

clusters the human homologues of which are found on chromosome 2 (Cazzaniga et al. 1994).

Structure of the XOR gene

Three sequences for the human XOR cDNA have been reported by different groups (Ichida

et al. 1993; Xu et al. 1994a; Saksela and Raivio 1996). The open reading frame of the human

XOR gene is composed of 3999 bp and accordingly encodes a protein of 1333 amino acids.

The cDNA reported by Saksela and Raivio (1996) codes for active XOR enzyme and is 99%

identical to the cDNA published by Ichida et al. (1993) and 94% identical to the sequence

reported by Xu et al. (1994a). Still another cDNA was initially reported to represent human


31

XOR cDNA, but was subsequently identified as the cDNA for AO based on the deduced

amino acid sequence and mRNA tissue distribution (Wright et al. 1993; Wright et al. 1995).

The amino acid sequence deduced from the human XOR cDNA reported by Ichida et al.

(1993) was 90% identical to the rat (Amaya et al. 1990) and 52% identical to the Drosophila

(Keith et al. 1987) XOR amino acid sequences. The deduced amino acid sequence of the rat

XOR cDNA (Amaya et al. 1990) was 94% identical to the deduced primary structure of the

mouse XOR protein (Terao et al. 1992).

The entire human XOR gene, spanning a DNA fragment of at least 60 kb, consists of 36

exons and 35 introns, ranging from 53 to 279 bp and approximately from 200 bp to at least 8

kb in size, respectively (Xu et al. 1996). The number of exons and the exon/intron junctions

of the human and mouse XOR genes have been conserved, whereas the third junction in the

rat gene differs from the other two (Cazzaniga et al. 1994; Chow et al. 1994; Xu et al. 1996).

Furthermore, the size of the second intron in the human and rat XOR genes exceeds 8 kb, but

is considerably shorter, 1.6 kb, in mouse. In Drosophila, the XOR gene comprises only four

exons, but shows striking homology to the mammalian XORs at the protein level (Keith et al.

1987; Ichida et al. 1993). In addition to the similarities in the structural and biochemical

properties of XOR and AO, the almost identical exon/intron structures of the genes coding for

these two enzymes suggest a common evolutionary origin (Terao et al. 1998; Garattini et al.

2003).

Regulatory regions of the XOR gene

Two transcriptional initiation sites, located –59 and –82 from the translational initiation

codon ATG, have been identified in the 5’-flanking sequence of the human XOR gene (Xu et

al. 1996). The function of the two separate potential transcriptional start sites is not known,

but mRNAs with different 5’-ends may facilitate post-transcriptional regulation or their

expression may be tissue-specific. Whereas in mouse only one transcriptional initiation site

was found, at least four transcriptional initiation sites have been recognized in rat, and their

correct usage has been shown to depend on the presence of the regulatory protein binding

sequences in the first exon (Cazzaniga et al. 1994; Chow et al. 1994; Chow et al. 1995; Clark

et al. 1998a; Clark et al. 1998b).


32

Approximately 2 kb of the human XOR promoter have been cloned and, based on the

promoter sequence, several potential transcription factor binding sites have been identified

(Xu et al. 1996). However, there are only a few reports on the function and the regulation of

the promoter for the human XOR gene (Xu et al. 2000). Consistent with the differences in the

levels of XOR tissue expression and activity between mouse and man, the levels of XOR

transcripts in mouse liver and the activity of a mouse promoter fragment (from 1 to 588

from the translational initiation codon) in transfection experiments were higher when

compared to human liver and to a corresponding human promoter fragment (from 1 to

463). A promoter region of the human XOR gene corresponding to the nucleotides from –1

to –138 from the translational initiation codon was shown to be necessary and sufficient for

basal promoter activity, whereas the region from –258 to –228 appeared critical for repressing

the core promoter activity (Xu et al. 2000). Results obtained by Xu et al. suggest that the

human XOR promoter carries a TATA-like element, which binds the components of the basal

transcription machinery (TFIID complex), including RNA polymerase II activity, and is

required for basal promoter activity. However, a consensus E-box, known to bind both

activators and repressors of transcription, was located at –240, and was assumed, together

with the TATA-like element binding protein, to restrict XOR promoter activity (Xu et al.

2000).

The promoter of the rat XOR gene has been cloned and the function of the proximal

promoter has been characterized (Chow et al. 1995; Clark et al. 1998a; Clark et al. 1998b).

Contrary to the human XOR promoter, no TATA-box was identified (Chow et al. 1994).

Instead, C/EBP proteins were shown to bind to the proximal promoter (Chow et al. 1995) and

together with transcription factor YY-1, which has the ability to bend DNA, to be necessary

for the basal promoter activity (Clark et al. 1998a). In addition, several other factors,

including NF-1, Oct-1, c-Myc and USF-related factors have been suggested to bind to the rat

XOR promoter (Clark et al. 1998b). The sequences extending to around –250 were shown to

elevate transcriptional activity of the rat promoter 50-fold compared to the promoterless

reporter gene construct. Lengthening of the promoter further up to –6000 decreased the

activity, indicating that repressive regulatory elements may exist (Chow et al. 1994).


33

Regulation of XOR under different oxygen levels and by nitric oxide

Oxygen sensing and hypoxia-inducible factor 1

The range of tissue and cell oxygen levels compatible with intact survival is relatively

narrow, and depends on the oxygen requirements of the specific tissue and cell type. The

main function of gene regulation by oxygen is to optimize the expression of proteins that help

cells to cope with either hypoxia or hyperoxia, and thereby to facilitate cell survival under

varying oxygen levels. Except for the characterization of hypoxia-inducible factor- (HIF- )

prolyl-hydroxylase (Ivan et al. 2001; Jaakkola et al. 2001; Safran and Kaelin 2003), the

mechanisms of oxygen sensing by eukaryotic cells are not well understood. Considering the

importance of oxygen homeostasis as a fundamental determinant of cellular energy

production and, ultimately, of cell survival, it is likely that several ways of oxygen sensing

and downstream signaling pathways have evolved. Heme and iron-sulfur clusters of proteins

have been suggested as possible sensors of changing oxygen levels (Semenza 1999a). Since

XOR is thought to contribute to the development of I-R injury, the regulation of gene

expression by oxygen is of special interest regarding the control of XOR expression.

HIFs are the best characterized mediators of hypoxic response (Semenza 1999b; Wenger

2002). HIF-1 is a heterodimeric transcription factor, which consists of - and -subunits and

recognizes the core recognition sequence 5’-RCGTG-3’ (Wang et al. 1995; Semenza et al.

1996). The HIF-1 -subunit has a basic helix-loop-helix DNA binding domain, and two other

HIF -subunits, namely HIF-2 and HIF-3 , have also been identified, but their precise role

has not been defined (Wenger 2002) The -subunit is identical to the heterodimerization

partner of the dioxin receptor/aryl hydrocarbon receptor, and is called aryl hydrocarbon

receptor nuclear translocator (ARNT). Furthermore, an inhibitory protein (IPAS), which can

bind to HIF-binding elements but lacks trans-activation capacity, has been recently

discovered (Makino et al. 2001). Interestingly, identification of a putative HIF-1 binding site

in the human XOR promoter has been reported (Hoidal et al. 1997).

HIF-1 regulates genes involved in processes leading to cellular adaptation and survival

under reduced oxygen tensions (Semenza 2000). Among HIF-1 target genes are

erythropoietin and vascular endothelial growth factor (VEGF) that support O2 delivery,


34

glycolytic enzymes and glucose transporters that participate in metabolic adaptation, and

transferrin, transferrin receptor and ceruloplasmin that affect cellular iron metabolism

(Semenza 1999b; Wenger 2002). In addition, several other genes with a variety of functions

have been identified as targets of HIF-1 (Wenger 2002).

In normoxic cells, HIF-1 is undetectable due to its degradation by a proteasome, but

under hypoxic conditions, the protein levels increase (Kallio et al. 1999). In cell culture, HIF-

1 protein levels and DNA binding activity begin to increase, when the ambient oxygen level

drops below 5%, and are maximal at 0.5% oxygen (Jiang et al. 1996). The mechanism of HIF-

1 degradation involves ubiquitination by the von Hippel-Lindau tumor suppressor protein

E3 ligase complex, which binds to the hydroxylated oxygen-dependent degradation domain of

HIF-1 . The hydroxylation of the critical prolines is catalyzed by the prolyl-hydroxylase,

which is dependent on O2 and iron as cofactors. The shortage of one or the other results in

HIF-1 protein stabilization due to diminished ubiquitination and lack of degradation of the

nonhydroxylated HIF-1 (Ivan et al. 2001; Jaakkola et al. 2001; Safran and Kaelin 2003). The

inhibition of the proteasome function alone does not result in transcriptional activation by

HIF-1, but HIF-1 phosphorylation, nuclear translocation, heterodimerization with ARNT,

DNA binding, and recruitment of general and tissue-specific transcription factors are required

for the transcriptional activation of the target genes (Wenger 2002). Any of the steps in HIF-1

activation may be modified by the oxygen supply. Still another level of regulation of HIF-1

activity is provided by recruitment of coactivators, including CPB/p300 (Kallio et al. 1998;

Wenger 2002). In addition to hypoxia, HIF-1 is induced by cytokines, growth factors and NO,

suggesting cross-talk with different signaling pathways (Semenza 2001; Semenza 2002;

Wenger 2002). Interactions with other DNA binding factors, for example activator protein-1

(AP-1), activating transcription factor-1 (ATF-1)/CREB-1, HNF-4, and nuclear factor B

(NF- B), are likely to be a prerequisite for tissue- and signal-specific activation of HIF-1.

Other mechanisms of regulation of gene expression by oxygen

Oxidative stress has been suggested to influence chromatin status by activation of histone

acetylases and inhibition of histone deacetylases, resulting in activation of genes required for

cellular adaptation and survival (Rahman 2002). On one hand, ROS produced by NADPH

oxidase have been proposed to contribute to HIF-1 inactivation in normoxia, whereas on the


35

other hand, ROS produced by mitochondria have been regarded as crucial contributors to the

activation of HIF-1 (Semenza 1999a; Michiels et al. 2002). In addition, many other

transcription factors are likely to be involved in gene regulation during hypoxia and

hyperoxia. For instance, the ubiquitous transcription factors Sp1 and Sp3 have been

implicated in the transcriptional activation of VEGF by oxidative stress (Schäfer et al. 2003),

whereas the induction of two glycolytic enzymes in hypoxia was mediated by down-

regulation of Sp3 (Discher et al. 1998). Putative binding sites for the redox-regulated

transcription factors NF- B and AP-1 have been identified in the promoter for the human

XOR gene (Xu et al. 1996).

NF- B plays a central role in the regulation of inflammatory and immune responses. It is

activated by cytokines, microbial products, and oxidative stress (Baeuerle and Henkel 1994;

Baldwin 1996; Barnes and Karin 1997; Michiels et al. 2002; Wang et al. 2002). In the lung

NF- B has been shown to be induced by acute hypoxia and contribute to I-R injury after

transplantation (Haddad 2003). NF- B has also been associated with endothelial dysfunction

in myocardial I-R injury (Boyle et al. 1999). NF- B exists as a homo- or heterodimer,

composed of members of the NF- B family including RelA (p65), RelB, c-Rel, p50 (NF-

B1), and p52 (NF- B2) (Baldwin 1996). The activation of NF- B is a multi-step process,

initiated in the cytoplasm and leading to nuclear translocation and DNA binding. ROS

produced in oxidative conditions activate NF- B, but to gain DNA binding activity reduced

conditions in the nucleus are required (Michiels et al. 2002).

AP-1 refers to dimeric transcription factors composed of Jun, Fos, or ATF subunits, which,

depending on the composition of the dimer, bind with varying efficacy either to AP-1 DNA

recognition elements or cAMP responsive elements (Karin et al. 1997; Shaulian and Karin

2001). AP-1 proteins are involved in growth control, transformation, inflammation and

immune response. Furthermore, a variety of environmental stresses, especially UV radiation,

induces AP-1 activity. Like NF- B, AP-1 is activated by ROS, but once in the nucleus,

reduction of its cysteine residues by nuclear protein Ref-1 and thioredoxin is required for

DNA binding (Hirota et al. 1997). Requirement of an AP-1 binding site for full transcriptional

activity conferred by HIF-1 in hypoxic regulation of VEGF suggests an additional link

between cellular oxygen levels and AP-1 (Damert et al. 1997).


36

Post-transcriptional regulation in hypoxia is exemplified by the stabilization of VEGF

mRNA by multiple proteins binding to its 3’-untranslated region (UTR) (Claffey et al. 1998,

Goldberg-Cohen et al. 2002). Expression of genes involved in iron metabolism may be

regulated by iron regulatory proteins (IRPs), which are RNA binding proteins that either

promote mRNA stability or inhibit protein translation (Eisenstein 2000). Hypoxia stabilizes

and increases RNA binding activity of IRP2 which, by binding to 3’-UTR in target mRNAs,

inhibits their degradation (Hanson et al. 1999). On the other hand, the RNA binding capacity

of IRP1 decreases during hypoxia, possibly due to the stabilization of its iron-sulfur [4Fe-4S]

cluster (Hanson and Leibold 1998).

Inhibition of HIF-1 degradation during hypoxia represents post-translational regulation by

oxygen. Interestingly, hypoxia has been shown to induce the accumulation of transcriptionally

active p53 by HIF-1 mediated protein stabilization (An et al. 1998), which may trigger

growth arrest or apoptosis (Carmeliet et al. 1998). Heat shock protein Hsp33, a potent

molecular chaperone, is activated by the formation of two disulfide bonds under oxidizing

conditions (Graf and Jakob 2002). Furthermore, proteins may be cross-linked by dityrosine

formation and transition metal containing proteins may be marked for proteolytic degradation

by oxidative modifications (Thannickal and Fanburg 2000). The post-translational regulation

of proteins by oxygen or ROS may represent the endpoint of signaling cascades initiated, not

always by oxygen, but for example by growth factors and cytokines.

Regulation of XOR in hypoxia

Induction of XOR activity during hypoxia has been proposed as one of the mechanisms

leading to the exacerbation of I-R injury by XOR. In rat lung XOR activity increased during

hypoxic exposure (Hassoun et al. 1998). Accumulation of XOR substrates, not the increment

of XOR activity, during ischemia was suggested to be responsible for the burst of free radical

generation by XOR upon reperfusion of rat myocardium (Xia and Zweier 1995). On the other

hand, consistent with the proposed role of elevated XOR activity in tissue injury caused by

hypoxia-reoxygenation, elevated XOR activity in hypoxia has been detected in several cell

lines (Terada et al. 1992; Hassoun et al. 1994; Lanzillo et al. 1996; Poss et al. 1996; Terada et

al. 1997; Kayyali et al. 2001). The level of XOR induction, however, remains controversial.


37

In bovine pulmonary artery endothelial cells total XOR activity showed a negative

correlation with oxygen concentrations ranging from 21% to 0%, with no change in the

relative amounts of XO and XDH. Production of O2 , attributed to elevated XOR levels,

increased after 48h anoxia followed by 4h reoxygenation, and was suggested to account for

increased oxidative cellular injury, neutrophil adherence, and albumin leakage (Terada et al.

1992). The effect of hypoxia (3% O2) to elevate XOR activity was confirmed in rat

epididymal fat pad endothelial cells, in which increased XOR mRNA levels, suggesting

transcriptional regulation, were detected (Hassoun et al. 1994). In support of transcriptional

induction, XOR mRNA levels were raised in rat pulmonary microvascular endothelial cells

exposed to 3% O2 (Lanzillo et al. 1996). On the contrary, XOR mRNA and protein levels

remained unchanged, despite increased enzyme activity, in bovine aortic endothelial cells

exposed to 3% oxygen for up to 48h (Poss et al. 1996). Interestingly, XOR activity increased

in mouse Swiss-3T3 cells cultured in 0% O2 for 24h by a post-translational mechanism, but

hypoxic exposure extended to 48h led to the elevation of XOR mRNA levels and protein

synthesis, implying pre-translational regulation (Terada et al. 1997). A mechanism for post-

translational regulation of XOR activity in hypoxia was suggested by a report indicating that

hypoxia-inducible protein kinase p38 and casein kinase II, involved in cell growth and

metabolism, phosphorylate XOR during hypoxia, thereby increasing enzyme activity in rat

pulmonary microvascular endothelial cells (Kayyali et al. 2001).

In conclusion, the variable and partly conflicting data on hypoxic induction of XOR may

be due to differences in exposure time and other experimental variables, but also to significant

differences in the overall XOR activity and tissue expression between species. Therefore,

assumptions directly applicable to the behavior of human XOR can not be made on the basis

of data obtained with animal cells.


38

Regulation of XOR in hyperoxia

Opposite to the induction in hypoxia, inhibition of XOR activity has been detected in

hyperoxia. XOR activity decreased in isolated rat lung as well as in bovine pulmonary artery

endothelial cells exposed to 100% or 95% O2, respectively, for 24h. In the same study,

purified XOR was inactivated by chemically produced ROS, by the addition of hypoxanthine

resulting in the generation of ROS by XOR itself, or by stimulated neutrophils, suggesting

that the inactivation may serve as a protective cellular control mechanism (Terada et al.

1988). As H2O2 is produced in the reaction catalysed by XO, it is interesting that the

generation of OH in the reaction of reduced XOR with H2O2 was implicated in XOR

inactivation (Terada et al. 1991). Whereas two studies indicated a concomitant reduction in

XOR activity and mRNA levels during exposure to 80% or 95% O2 (Hassoun et al. 1994;

Lanzillo et al. 1996), another study did not reveal changes in either XOR activity or mRNA in

cells cultured first for 24h in hypoxia and subsequently for 24h in normoxia (21% O2) (Poss et

al. 1996). Consistent with the inactivation of purified XOR by ROS, increased extracellular

H2O2 production decreased XOR activity in bovine pulmonary artery endothelial cells

(Hassoun et al. 1995).

Effects of nitric oxide on XOR activity

Even though inflammatory cytokines are thought to induce XOR activity, it was clearly

diminished in rat and mouse macrophages after interferon- stimulation, despite increasing

mRNA levels. Based on the effects of a nitric oxide (NO) inhibitor, the induced production of

NO was suggested to be the cause of XOR inactivation and to represent a protective

mechanism against XOR-mediated tissue injury (Rinaldo et al. 1994). In another study,

exposure of cells directly to NO or NO-generating agents reversibly inhibited XOR activity

without changing its mRNA levels (Hassoun et al. 1995). The results indicating XOR

inactivation by NO were later argued, and it was suggested that in biological systems, due to

presence of more readily oxidized molecules than XOR, inactivation would not occur

(Houston et al. 1998).

However, NO was shown to inhibit the activity of purified bovine milk XOR, by

inactivating reduced XO and XDH. Based on electron paramagnetic resonance studies,

preservation of the activities of the iron-sulfur and FAD centers, and reactivation of the NO


39

inactivated enzyme by a sulfide-generating system, NO was suggested to react with the

critical sulfur atom in the Mo center leading to the conversion of XO and XDH into the

inactive desulfo forms (Ichimori et al. 1999). Another study demonstrated that NO itself

would not inactivate XO, but, in the presence of O2 , ONOO would be generated leading to

the inactivation of XO (Lee et al. 2000). Yet, as discussed above, there are limitations to the

applicability of these results conducted in vitro with purified XOR to living cells.

Interestingly, O2 generated by XOR was shown to react with NO and inhibit NO-dependent

vascular relaxation in patients and mice with sickle cell disease. In this condition increased

plasma XO levels were thought to derive from the liver experiencing repeated episodes of

hypoxia-reoxygenation (Aslan et al. 2001).

Regulation of gene expression by iron

Iron and ischemia-reperfusion injury

Despite the fundamental importance of iron for normal cellular functions, elevated levels

of free iron are detrimental to cells, since iron may promote the formation of the extremely

reactive OH from other ROS in Haber-Weiss or Fenton reactions, causing protein oxidation,

lipid peroxidation, and DNA damage (Halliwell 1987; Halliwell and Gutteridge 1989; Chan

1996; Davalos et al. 2000). Ischemia has been shown to lead to iron deposition in rat brain

(Chi et al. 2000). Furthermore, high plasma and central nervous fluid ferritin concentrations,

reflecting body iron stores, have been associated with stroke progression in patients with

acute cerebral infarction (Davalos et al. 2000). Iron deposition in brain may be a marker of

neuronal damage, since it may reflect the accumulation of iron in macrophages and microglia

(Chi et al. 2000). Alternatively, acidosis in ischemic tissues may lead to dissociation of iron

from storage molecules, and as a result, increase in free iron may exacerbate tissue damage

(Siesjö et al. 1985). In the isolated rabbit lung, prolonged ischemia resulted in release of iron

into the vascular space (Huang et al. 2001).

It has been suggested that a relationship exists between elevated iron levels and

cardiovascular disease. An unfavourable modification of low density lipoproteins has been

proposed as one of the mechanisms (de Valk and Marx 1999). On the other hand, iron


40

chelation by desferrioxamine (DFO), has been shown to improve NO-mediated vasodilation

in patients with coronary artery heart disease (Duffy et al. 2001). Interestingly, DFO was

shown to decrease XOR activity in cell culture (Rinaldo and Gorry 1990) and the rat XOR

mRNA carries a potential 5’-iron responsive element (IRE) conserved in the human 5’-UTR

(Chow et al. 1995). Considering the proposed role of XOR in the pathogenesis of I-R injury,

the regulation of XOR by iron is of interest.

Post-transcriptional regulation by iron

Due to the absolute requirement of iron and, on the other hand, the potentially detrimental

effects of excess iron, cellular iron homeostasis is carefully controlled by iron absorption in

the intestine, by cellular iron intake, and by iron storage. Furthermore, at the molecular level

the expression of proteins involved in iron metabolism is regulated post-transcriptionally by

IRPs. IRP1 and IRP2 promote the stability of target mRNAs by binding to 3’-UTR IREs or

inhibit protein translation by binding to IREs in the 5’-UTR. During iron depletion, IRPs bind

to the 3’-IREs of transferrin receptor mRNA and retard its degradation. As a result, the

expression of the receptor and eventually cellular intake of iron increase. Simultaneously,

binding of IRPs to the 5’-IRE on the mRNA of ferritin, the main iron storage protein, inhibits

its translation. In iron-replete cells, a [4Fe-4S] iron-sulfur cluster is assembled in IRP1

converting it into cytosolic aconitase and inhibiting its RNA binding capacity. On the other

hand, iron has been suggested to induce oxidation of IRP2, leading to its proteasomal

degradation (Eisenstein 2000; Templeton and Liu 2003). In addition to cellular iron

concentration, IRP activities are influenced by oxygen levels (Hanson and Leibold 1998;

Hanson et al. 1999).

Transcriptional regulation by iron

Not only genes involved in iron metabolism are responsive to iron, but also genes related

to oxygen, oxidative stress, energy metabolism, cell cycle regulation, and tissue fibrosis are

regulated by iron, not only by IRP-mediated, but also by transcriptional mechanisms

(Templeton and Liu 2003). Iron chelation has been shown to induce HIF-1 activity, because

iron is essential for the activity of the prolyl-hydroxylase required for HIF-1 protein

degradation by the proteasome (Ivan et al. 2001; Jaakkola et al. 2001). Consequently,

reduction in cellular iron may lead to increased levels of HIF-1 and transcriptional induction


41

of the target genes. The transcriptional regulation of transferrin receptor by iron chelation

probably involves HIF-1 (Bianchi et al. 1999). Other examples of genes transcriptionally

regulated by iron include protein kinase C, cyclin-dependent kinase inhibitor p21,

retinoblastoma susceptibility protein pRb, many stress proteins, and inducible NO synthase,

but also ferritin and transferrin are regulated at the level of transcription by elevated iron

levels or iron depletion, respectively (Alcantara et al. 1994; Dlaska and Weiss 1999; Ye and

Connor 2000; Alcantara et al. 2001; Templeton and Liu 2003). Iron-induced transcription

may result from signaling through increased intracellular production of ROS that activates

transcription factors like NF- B, or from direct lipid peroxidation (Brenneisen et al. 1998;

Michiels et al. 2002; Templeton and Liu 2003). However, it is likely that many more

transcription factors are affected by iron and oxidative stress. Currently, the precise

mechanisms of transcriptional regulation by iron are unclear.

Aims of the Study

42

AIMS OF THE STUDY

Based on the hypothesis that XOR has a role in the pathogenesis of I-R injury, we have

investigated factors regulating XOR expression. We aimed at functional characterization of

the human XOR promoter, identification of transcription factors affecting XOR expression,

and understanding the regulation of XOR expression under specific circumstances related to

I-R injury.

The specific aims of this study were:

1) To determine the chromosomal localization of the human XOR gene,

2) to characterize the function of the proximal human XOR promoter and identify

transcription factors regulating XOR gene transcription,

3) to study XOR expression and its regulation during the small-intestinal enterocyte-like

differentiation of an intestinal cell line,

4) to study the effect of different oxygen levels on XOR activity and gene transcription, and

5) to investigate the effect of varying intracellular iron levels on XOR expression.

Materials and Methods

43

MATERIALS AND METHODS

Isolation of XOR-specific clones

Screening of DNA libraries

To obtain a human XOR cDNA clone, a 249 bp cDNA fragment, corresponding to the

human XOR cDNA nucleotides from 3550 to 3798, was isolated from human breast cDNA

library (Clontech, Palo Alto, CA, USA) by polymerase chain reaction (PCR) using the

primers 5’-GGTTCCAGCTTGAATCCTGCCATTGATATTGGACAA-3’ (forward) and 5’-

GAAAAGAGGTGGCTCCCCCAACAGCCTTGGA-3’ (reverse), designed on the basis of

homology between Drosophila (Keith et al. 1987) and rat (Amaya et al. 1990) XOR cDNAs.

The PCR fragment was cloned into the TA cloning vector (Invitrogen, Groningen, The

Netherlands) and, subsequently, a RNA probe was created and used to screen a human breast

cDNA library (Clontech), as well as a human lymphocyte genomic cosmid library (Clontech).

A 2 kb cDNA clone and an approximately 20 kb genomic clone were isolated. The genomic

clone was characterized by southern blot analysis using probes corresponding to the original

249 bp cDNA fragment and to the 2 kb cDNA, and partially sequenced.

Cloning of human XOR gene promoter fragments

Human XOR gene promoter fragments XOR1 and XOR2 were isolated by PCR from

human genomic DNA, and subsequently XOR4, XOR5, and XOR6 were generated by using

XOR2 as the template. The primers were designed on the basis of the sequence of the human

XOR gene promoter (Xu et al. 1996) and included restriction enzyme sites for BglII or MluI

in their 5’-ends and for HindIII in 3’-ends. The 5’-primer sequences were

5’-GGGAAGATCTTGTGGTTTGTAGGATGTTTAGT-3’ (complementary to nucleotides

1937 to 1913 in the human sequence, underlined), 5’-GGGGACGCGTCTTACTTAAGGA

AGGCTGGC-3’ ( 1174 to –1153), 5’-GTGGAGATCTTAATTTGCTGTGTGTGATTGTT-

3’ ( 224 to 203), 5’- CTGAAGATCTGAGCTGGTTCCCTCCCATTG-3’ ( 142 to 120),

and 5’-TGGGACGCGTAACTTTCAGGTCACAGAGCA-3’ ( 92 to 69) corresponding to

promoter fragments in XOR1, XOR2, XOR4, XOR5 and XOR6, respectively. The 3’-primer

sequence for all PCR reactions was 5’-CTACTAAGCTTTCTGCCATTCACAAAGAAAAC-

3’ (+42 to +19). XOR3 was obtained by shortening XOR1 by exonuclease III digestion

(Erase-a-Base System, Promega, Madison, WI), and its sequence corresponds to nucleotides


44

547 to +42. XOR5mut, with mutated inverted CCAAT-box, was created by PCR using

XOR2 as the template. The mutagenic primer was 5’-CTGAAGATCTGAGCTGGTTCCC

TCCCATCAGTGGACCT-3’ (mutated base pairs underlined), the 5’-end of the primer

containing a BglII site. DNA fragments XOR6Emut1 and XOR6Emut2, with mutations in the

GATA consensus sequence, were prepared by PCR using XOR5 as the template, and the

mutagenic 5’-primers were 5’-TGGGACGCGTAACTTTCAGGTCACAGAGCAGTCAT-3’

and 5’-GGGACGCGTAACTTTCAGGTCACAGAGCAGCCCTA-3’ (mutated base pairs

underlined), respectively. The 3’-primer was as above. The PCR products and XOR3 were

cloned into pGL3-basic (Promega) bearing a luciferase gene and sequenced.

Somatic cell hybrid mapping panel

The chromosomal localization of the human XOR gene was initially studied by PCR

performed with DNA extracted from human × mouse and human × hamster cell hybrids

obtained from the NIGMS Human Genetic Mutant Cell Repository (Somatic Cell Hybrid

Mapping Panel No. 1). Primers used in PCR were designed according to the sequences at the

5’- and 3’-ends of a 0.7 kb EcoRI-fragment of the cosmid clone, which hybridized with a

probe corresponding to the 2 kb cDNA clone described above. A PCR product was obtained

with human, but not with any rodent, genomic DNA as the template.

Polymerase chain reaction (PCR)

DNA (50 ng) from each cell hybrid, 50 100 ng genomic DNA, or 100 ng of XOR2 or

XOR5 were used as templates in PCR. Taq (Promega) or Dynazyme (Finnzymes, Espoo,

Finland) DNA polymerases were used in all reactions, except for the generation of XOR1,

where Expand High Fidelity DNA polymerase (Roche Molecular Biochemicals, Indianapolis,

IN, USA) was utilized.

Fluorescence in situ hybridization (FISH)

To determine the chromosomal localization of the human XOR gene, FISH with a biotin-

11-dUTP-labeled probe (Nick Translation Kit, Bethesda Research Laboratories, Bethesda,

MD, USA) was performed essentially as described (Pinkel et al. 1986; Lichter et al. 1988).

On metaphase spreads, made on microscope slides from phytohemagglutinin-stimulated

normal lymphocytes by standard procedures, 50 ng of the labeled 20 kb genomic XOR


45

cosmid probe or 300 ng of the 2 kb XOR cDNA probe were applied. Subsequently, the

hybridized probe was detected using fluorescein isothiocyanate (FITC) -avidin (Vector

Laboratories, Burlingame, CA, USA) and amplified once. Nuclear DNA was counter-stained

with propidium iodide and stained with 4’-6’-diamino-2-phenylindole (DAPI). The slides

were examined with a microscope and photographed. Digitized image analysis was performed

using Nikon SA fluorescence microscope, a high-resolution CCD camera (Xillix, Richmond,

Canada), and software as described (Kallioniemi et al. 1994). Eight two-color images of the

DAPI counterstain and FITC hybridization signal were collected, overlaid electronically, and

analyzed for the signal location as well as for the fractional length from the terminus of the

short arm (Flpter), as described (Kallioniemi et al. 1994).

Cell culture and exposures of cells

Standard cell culture

Human embryonic kidney 293T (a kind gift from Prof. Kalle Saksela, University of

Tampere, Finland), mouse fibroblast NIH-3T3, and human colon carcinoma Caco-2 cells

(ATCC, Manassas, VA, USA) were grown in Dulbecco’s modified Eagle’s medium with

Glutamax (Gibco, Paisley, UK) supplemented with 10% (v/v) fetal calf serum or, for NIH-

3T3 cells, with calf serum, 50 100 U/ml penicillin, and 50 100 g/ml streptomycin in 5%

CO2 at 37 C. BEAS-2B SV-40 transformed normal human bronchial epithelial cells (ATCC)

were cultured in serum-free, hormone-supplemented bronchial epithelial cell growth medium

(Cytotech ApS, Hellebaek, Denmark).

Caco-2 cell differentation

For in vitro differentiation, Caco-2 cells were seeded at 5 x 105 cells/10-cm cell culture

dish, medium was changed one day after plating and every 2 3 days thereafter.

Hypoxia, hyperoxia, and cobalt chloride

Confluent BEAS-2B cells were exposed to either 0.5%, 3%, 21%, or 95% O2 for 4 48 h.

Fresh medium containing 75 M CoCl2 was applied, and the cells incubated at 21% O2 for 24

h. Transfected 293T cells, carrying XOR promoter constructs or hypoxia-responsive reporter

construct (HRE-luc), were exposed to 0.5% or 21% O2 for 24 h. The specified O2


46

concentrations were achieved by infusing a preanalyzed gas mixture into airtight chambers

(Billups-Rothenburg, Del Mar, CA, USA).

Iron, copper, iron chelators, and hydroxyl radical scavengers

Concentrations of ferric ammonium citrate (FAC) (Riedel-de-Haen, Seelze, Germany)

between 180 M to 1.8 mM; 1 mM FeSO4, 10 to 100 M CuSO4, and 30 mM dimethyl

sulfoxide (DMSO) (all three from Merck, Darmstadt, Germany); and 100 M DFO, 10 mM

1,3-dimethyl-2-thiourea (DMTU), 1 mM N-acetylcysteine (NAC), and 5 M 1,10-

phenanthroline (Sigma, St. Louis, MO, USA) were applied on subconfluent cells for 6 24 h.

Protein and RNA synthesis inhibitors

To inhibit protein synthesis, cells were cultured with 10 g/ml cycloheximide (Sigma) for

6 24h. Correspondingly, RNA synthesis was inhibited with 1 g/ml actinomycin D (Sigma).

Transfections and reporter gene analysis

Transfections with XOR promoter constructs and HRE-luc

For functional XOR promoter analysis, 293T cells were seeded on 12-well plates (2 x 105

cells per well) and NIH-3T3 cells on 6-well plates (3 x 105 per well) 24 h before transfection,

and transfected with 0.5 g or 1 g of XOR promoter constructs, respectively, using FuGene6

(Roche Molecular Biochemicals) transfection reagent. In all experiments, 10 ng (293T) or 0.5

g (NIH-3T3) of pCMV (Clontech), a -galactosidase expression vector, were cotransfected

to monitor transfection efficiency, and empty pGL3-basic vector was used as a control. To

study the response of the XOR promoter constructs in hypoxia, 293T cells were seeded onto

12- or 6-well plates (2 x 105 or 5 x 105 cells per well, respectively) and transfected with 0.33

or 1 g of XOR promoter constructs. HRE-luc carrying three tandem copies of the

erythropoietin hypoxia responsive element coupled to luciferase was kindly provided by Dr.

Pekka Kallio (Kallio et al. 1998), and used as described for the XOR promoter constructs. In

all transfection experiments, standard culture medium or medium containing 1 or 2 mM FAC,

was changed 16 h after transfection and the cells were further incubated for 24 h in 21% or

0.5% O2. Luciferase activity was determined with reagents from Promega using a

Luminoskan RT reader (Labsystems, Helsinki, Finland). -galactosidase activity was

determined according to Rosenthal (1987).


47

Expression of XOR protein and GATA-4

When reaching about 50% confluence, 293T cells in 10-cm cell culture dishes were

transiently transfected, using FuGene6, with 2 g of pcDNA3-expression vector (Invitrogen)

carrying the complete coding sequence of human XOR (Saksela and Raivio 1996) named

pcDNA3-XOR, and after 17 h fresh medium with 180 M or 1 mM FAC or 100 M DFO

was added, followed by 24 h incubation. To prepare nuclear extracts rich in GATA-4 protein,

293T cells were transiently transfected with 10 g of GATA-4 expression plasmid pMT2-

GATA-4 (Arceci et al. 1993).

Assays of enzyme activities and iron measurement

XOR activity

For the activity measurements cells were washed twice or three times with PBS and

harvested in 50 mM potassium phosphate buffer, pH 7.8, containing 0.5 mM dithiothreitol, 1

mM EDTA, 0.5 g/ml leupeptin, and 0.2 mM PMSF, or 50 l/ml protease inhibitor cocktail

(Sigma). Subsequently, the cell suspension was sonicated on ice or frozen and thawed twice

and centrifuged at 4 C at 15 800 g for 8 min. XOR activity of the supernatant was expressed

as nmol/min/mg protein.

Total XOR and XO activities from cells were determined using 40 M 14C-xanthine

(specific radioactivity 55 60 mCi/mmol) (NEN Life Science Products, Boston, MA, USA) as

substrate in the presence or absence of 400 M NAD+, respectively. After incubation at 37 C,

the reaction was stopped by adding perchloric acid and subsequently neutralized with

potassium hydroxide. The product uric acid was separated by HPLC (Shimadzu, Kyoto,

Japan) with a reverse-phase column and eluted with 50 mM potassium phosphate, pH 4.5,

followed by quantification with Radiomatic Flow Scintillation Analyzer (Packard Instrument,

Meriden, CT, USA).

The effect of 30 or 90 mM H2O2 on the activity of purified bovine milk XOR (Biozyme

Laboratories, South Wales, UK) was measured spectrophotometrically by detecting

absorbance change at 295 nm, corresponding to uric acid production, over 3 min at the start

and at the end of 30 min incubation.


48

Analysis of 2,6-dichlorophenolindophenol reduction and NADH oxidation

The electron transfer activity of purified XOR from xanthine to the artificial substrate 2,6-

dichlorophenolindophenol (DCPIP) was determined spectrophotometrically by measuring the

absorbance of DCPIP at 600 nm. NADH oxidation of cell lysates was measured by

monitoring the absorbance of NADH at 340 nm.

Maltase activity

Maltase activity was measured according to the method by Messer and Dahlqvist (Messer

and Dahlqvist 1966). To prepare cell homogenates for the assay, Caco-2 cells were washed

three times with PBS and mechanically harvested in saline. The cell suspensions were

homogenized for 45 s, followed by centrifugation at 15 800 g for 40 50 min at 4 C, and the

supernatant was stored at 80 C.

Intracellular iron measurement

Cells treated with iron or iron chelators were washed three times with PBS, harvested in

PBS, frozen and thawed twice, and centrifuged at 4 C 15800 g for 8 min. The iron content of

the supernatant was measured by using Rauta Kit 141010 (Reagena Ltd, Kuopio, Finland), in

which iron is first released in denaturing conditions (pH 4.8), and then reduced by ascorbic

acid to ferrous iron, which forms a stable complex with the chromogen ferene-S. This

complex is then measured spectrophotometrically at 595 nm.

Protein analysis

Protein quantification

Total protein concentrations were determined using Bio-Rad DC protein assay (Bio-Rad

Laboratories, Hercules, CA, USA) and XOR protein was quantified by ELISA as described

(Sarnesto et al. 1996).

Western blot analysis

Proteins from NIH-3T3 (10 g) or 293T cells overexpressing XOR (2.5 g), harvested as

for the activity measurements, were separated by 7.5% sodium dodecyl sulphate-

polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to Immobilon-P membrane

(Millipore, Bedford, MA, USA). The blotted membrane was blocked with 5% (w/v) skim


49

milk in 0.1 M Tris, 1 M NaCl and 0.1% (v/v) Tween-20 for 1 h. For detection of XOR

protein, polyclonal human anti-XOR antibodies (Sarnesto et al. 1996) at a dilution of 1:1000

(for NIH-3T3 proteins) or 1:2000 (for cells overexpressing XOR) were used, followed by

horseradish peroxidase-conjugated goat anti-rabbit secondary antibody (Jackson Immuno-

Research Laboratories, West Grove, PA, USA) at a dilution of 1:5000. To control for the

protein loading, monoclonal anti- -tubulin antibody (Sigma) at a dilution of 1:40000 was

applied on the same Western blot. The protein bands were visualized by enhanced

chemiluminescence (Amersham Pharmacia Biotech, Buckinghamshire, UK).

To detect NF-YA, GATA-4, or GATA-6, nuclear proteins (20 g), were separated by 10%

SDS-PAGE. Rabbit polyclonal anti-NF-YA antibody (Chemicon, Temecula, CA, USA) was

used at a dilution of 1:1000, followed by secondary antibody as above. Goat polyclonal anti-

GATA-4 and -GATA-6 antibodies (both from Santa Cruz, Santa Cruz, CA, USA) were used

at dilutions of 1:500, and horseradish peroxidase-conjugated rabbit anti-goat secondary

antibody (Dako, Glostrup, Denmark) at a dilution of 1:2000. To control for the protein

loading, mouse monoclonal anti- -tubulin antibody (Sigma) was applied at a dilution of 1:50

000.

Immunoprecipitation

Proteins (0.8 mg) from Caco-2 cell extracts were diluted 1:2 with PBS containing 1%

nonidet P-40 (v/v), 1 mM EDTA, and 50 l/ml protease inhibitor cocktail (Sigma) (IP-buffer)

and incubated with 3 l of polyclonal human anti-XOR antibodies for 90 min at 4 C.

Subsequently, 50 l protein G sepharose (Amersham Biosciences, Uppsala, Sweden) were

added, and incubation continued for further 90 minutes. The samples were washed four times

with IP-buffer, bound proteins eluted with reducing Laemmli-sample buffer, separated and

detected as described for the Western blot analysis of XOR.

RNA analysis

Northern blot analysis

Total RNA was extracted using RNeasy Mini Kit (Qiagen, Hilden, Germany) and

separated on a 1% agarose-formaldehyde gel for Northern blot analysis. RNAs were

transferred to Biodyne A nylon filters (PALL Gelman Sciences, Portsmouth, UK) and fixed


50

by UV cross-linking. To detect XOR mRNA, membranes were hybridized using standard

procedures (Sambrook et al. 1989) with [32P]-labeled complementary RNA probe

corresponding to nucleotides 321-3021 of the rat XOR cDNA (kindly provided by Professor

T. Nishino, Nippon Medical School, Tokyo, Japan) (Amaya et al. 1990). The same

membranes or membranes containing RNA from Caco-2 cells were reprobed with random

primed (Prime-a-Gene Labeling System, Promega) mouse 18S ribosomal gene DNA probe

(Ambion, Austin, TX, USA) to control for RNA loading. The autoradiography films were

scanned and analysed with the Scion Image beta 4.0.2 analysis software (Scion Corporation,

Frederick, MD, USA).

Ribonuclease protection assay (RPA)

RNA from Caco-2 cells was extracted as above. Specific RNA was quantified using RPA

according to the manufacturer’s protocol (RPAIII, Ambion). [32P]-labeled antisense RNA

probe was transcribed from a DNA template corresponding to the nucleotides 405-789 of the

human XOR cDNA (Saksela and Raivio 1996), and 100 000 cpm of the radiolabeled probe

were hybridized with 30 g of RNA overnight at 42 C. To control for the RNA content, 30

000 cpm of a RNA probe transcribed from human -actin cDNA (pTRI-Actin-Human,

Ambion) were added to the same hybridization reaction with the XOR probe. After RNase

digestion, the protected fragments were separated by 5% polyacrylamide, 8 M urea gel, which

was exposed to autoradiography films, scanned, and analyzed as described above.

RNA from BEAS-2B cells was extracted by acid guanidium thiocyanate-phenol-

chloroform extraction method according to Chomczynski and Sacchi (1987). RPA was

performed essentially as above, but 20 g of RNA were hybridized with 60 000 cpm of the

XOR probe, and -actin was detected from separate hybridization in a parallel assay.

Nuclear protein extracts

Cells were harvested in PBS and nuclear protein extracted as described (Andrews and

Faller 1991). After lysis in a buffer containing 10 mM Hepes, pH 7.9, 1.5 mM MgCl2, 10 mM

KCl, 1 mM dithiotreitol, 0.5 mM PMSF, 3 g/ml leupeptin and 5 g/ml pepstatin, the cells

were left on ice for 15 min and then vortexed for 10 s. After centrifugation for 5 s at 15 800 g,

proteins from the nuclear pellet were extracted for 20 min on ice in a buffer (20 l/nuclei


51

from 107 cells) containing 20 mM Hepes, pH 7.9, 25% (vol/vol) glycerol, 1.5 mM MgCl2, 0.2

mM EDTA, 420 mM NaCl, 1mM dithiotreitol, 0.5 mM PMSF, 3 g/ml leupeptin and 5 g/ml

pepstatin. Nuclear debris was removed by centrifugation for 5 min at 4 C at 15 800 g.

Electrophoretic mobility shift assay (EMSA)

Probes

Electrophoretic mobility shift assay (EMSA) probes XOR5E and XOR5Emut were created

by digesting the plasmids XOR5 and XOR5mut with BglII and HindIII. Probes XOR6E,

XOR6Emut1, and XOR6Emut2 were prepared by digestion of the plasmid XOR6 and the

plasmids carrying XOR6Emut1 and XOR6Emut2 with MluI and HindIII. The restriction

fragments were purified using 8% PAGE. Oligo1 (nucleotides –135 to –107) was prepared by

annealing the oligonucleotides 5’-GAGCTGGTTCCCTCCCATTGGTGGACCTTATTT-3’

and 5’-AATGAAATAAGGTCCACCAATGGGAGGGAACCA-3’ and oligo2 (nucleotides

92 to 60) by annealing the oligonucleotides 5’-GCGTAACTTTCAGGTCACAGAGC

AGTGATAACT-3’ and 5’-AGTTATCACTGCTCTGTGACCTGAAAGTTACGC-3’. NF-Y,

C/EBP, and GATA consensus, and the mutated C/EBP double-stranded oligomers used in

competition assays were purchased from Santa Cruz. The probes were 5’-end labeled with -32P ATP using T4 polynucleotide kinase (Promega). EMSA probes, except for the

commercial oligomers, are presented in Table 2. Oligo1 and oligo2 are also shown in Figure

7.

Table 2. Probes used in EMSAs.

Probe Nucleotides Regulatoryelement

Sequence

XOR5E 142 to +42 inverted CCAAT 5’…CCCTCCCATTGGTGGACC…3’

XOR5Emut 142 to +42 inverted CCAAT 5’…CCCTCCCATCAGTGGACC…3’

XOR6E 92 to +42 GATA 5’…AGAGCAGTGATAACTACCTG…3’

XOR6Emut1 92 to +42 GATA 5’…AGAGCAGTCATAACTACCTG…3’

XOR6Emut2 92 to +42 GATA 5’…AGAGCAGCCCTAACTACCTG…3’

oligo1 135 to 107 inverted CCAAT 5’-GAGCTGGTTCCCTCCCATTGGTGGACCTTATTT-3’

oligo2 92 to 60 GATA 5’-GCGTAACTTTCAGGTCACAGAGCAGTGATAACT-3’

Regulatory elements are shown in bold, mutated nucleotides underlined.


52

Binding reactions

Nuclear proteins were incubated on ice for 10 min with 2 g poly(dI-dC)(dI-dC)

(Amersham Pharmacia Biotech) in 20 mM Hepes (pH 7.9), 10% glycerol (vol/vol), 50 mM

KCl, 0.5 mM EDTA, 1 mM DDT, 1 mM MgCl2, 0.5 mM PMSF and 1 M leupeptin. A 5’-

end-labelled probe (5000–10000 cpm) was then added and the incubation was continued at

22 C for 30 min. In competition experiments, 10 100-fold molar excess of the unlabeled

probe was added before the labeled probe. In supershift experiments, 2 g of anti-NF-YA,

anti-NF-YB or anti-C/EBP- antibodies and 1 g of anti-GATA-4 or -GATA-6 antibodies

were added after binding reactions and incubation was further continued for 50 min at 22 C.

Except for the anti-NF-YA antibodies (Chemicon), the antibodies for supershift assays were

from Santa Cruz. Reaction products were separated by 4% polyacrylamide gels run in 22.5

mM Tris-borate, 0.5 mM EDTA for 2 3 h at 200 V at 22 C. After electrophoresis, the gels

were dried and visualized by autoradiography.

Statistics

Results are expressed as means SD, and means were compared by using two-tailed t-test

with unequal variations. Values of p<0.05 were considered significant.

Results

53

RESULTS

Localization of the human gene for XOR on chromosome 2p22 (I)

As an initial step for the characterization of the human XOR gene, we attempted to

determine the chromosomal localization of the gene, which had tentatively been assigned to

chromosome 2 (Ichida et al. 1993). Human XOR cDNA and genomic clones were first

isolated, a human × rodent hybrid cell panel analyzed by PCR with human specific primers,

and finally FISH was performed.

In the analysis of the human × rodent hybrid cell panel, the 0.7 kb PCR product specific

for human genomic DNA was obtained in three out of seven hybrid cell samples containing

human chromosome 2, whereas no amplification product was visible in the 16 samples

lacking human chromosome 2. Based on this the human gene for XOR was initially localized

to chromosome 2.

For more precise localization on chromosome 2, FISH was performed. The hybridization

of the fluorescent 20 kb genomic cosmid probe on 174 lymphocyte metaphase spreads in two

separate experiments was analyzed. In 159 (91%) metaphases fluorescent signals were

detected on the short arm of chromosome 2 (2p), and 66 (38%) metaphases exhibited

symmetrical signals on both arms of the chromosome. In 131 (75%) metaphases signals were

detectable only on chromosome 2, whereas 38 (22%) showed nonspecific binding. The 2 kb

cDNA probe hybridized with chromosome 2 on 28 (34%) out of 82 samples without

background, and no signal was seen on 39 (48%) metaphase spreads. Analysis of the digitized

images of the hybridization signal with the cosmid probe overlayed on DAPI images of the

corresponding samples indicated a chromosomal localization of the human XOR gene on

bands 2p22.3 p22.2 and a FLpter value of 0.135 0.0164.

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54

Transcription factor NF-Y regulates the human XOR gene promoter (II)

At the outset of these studies, the function of the proximal promoter of the rat XOR gene

had been characterized (Chow et al. 1994, Chow et al. 1995; Clark et al. 1998a, Clark et al.

1998b), but no functional data were available on the regulation of the human XOR gene.

However, 2 kb of the 5’-UTR had been cloned, and based on its sequence, several putative

cis-regulatory elements had been identified (Xu et al. 1996). The 5’-UTR sequences of the

human and rat genes showed considerable differences, suggesting different transcriptional

regulation. We therefore decided to clone and characterize the function of the human XOR

promoter.

Functional characterization of the human XOR promoter

A human XOR promoter fragment, corresponding to nucleotides –1937 to +42 relative to

the translational initiation site, was obtained by PCR from human genomic DNA and named

as XOR1. Subsequently, a 5’ deletion series of XOR promoter constructs was prepared. The

sequence obtained by sequencing both strands of XOR1 was submitted to GenBank

(GenBank AF203979). XOR5 (from –142 to +42) showed highest promoter activity in

transiently transfected human 293T cells (II, Fig. 1A), whereas XOR1 was most active in

NIH-3T3 cells (II, Fig. 1B). Based on the functional data, the sequence lacking from XOR6

(from –92 to +42), but included in XOR5 was hypothesized to carry positive regulatory

elements. Two inverted CCAAT-boxes (from 123 to –119 and from –100 to –96) (II, Fig.

1C) were identified. The one spanning from 123 to –119 and partially overlapping the

element corresponding to rat C/EBP binding site was subsequently mutated and XOR5mut

was generated to be able to evaluate the importance of this cis-element for the promoter

activity. The mutation of the CCAAT-box led to a 45% reduction in relative luciferase

activity compared to the construct with intact sequence, suggesting an important functional

role for this element in the transcriptional regulation of the human XOR gene.

NF-Y binds to the proximal human XOR promoter, but C/EBP- does not

Oligo1, corresponding to the XOR promoter region from –135 to –107 and carrying the 5’

inverted CCAAT-box, and XOR5E, corresponding to the promoter construct XOR5, both

showed binding of proteins in EMSAs performed with nuclear extracts from 293T and NIH-

Results

55

3T3 cells. One of the bands formed with XOR5E as the probe was competed with unlabeled

oligo1 and NF-Y consensus oligonucleotide and, additionally, a supershift was shown with

NF-YB antibody. Similarly, the major band formed with oligo1 was competed with NF-Y

consensus oligonucleotide and recognized by NF-YB antibody, whereas binding of proteins to

oligo1 was not influenced by XOR5Emut with the mutated CCAAT-box, consensus or

mutated C/EBP oligonucleotides or C/EBP- antibody. Taken together, our data indicate that

the proximal human XOR promoter carries a functionally important NF-Y binding site, but

does not bind C/EBP proteins.

Transcriptional induction of XOR during enterocytic differentiation of

Caco-2 colon carcinoma cells – A role for NF-Y (III)

High XOR activity and protein levels have been detected in the human small intestine

(Sarnesto et al. 1996; Linder et al. 1999). Inhibition of XOR activity has been associated with

the alleviation of experimental intestinal I-R injury (Parks et al. 1982; Albuquerque et al.

2002), whereas the addition of exogenous xanthine augmented apoptotic tissue injury

(Genesca et al. 2002). Factors regulating XOR expression in the intestine have not been

defined. Considering our results on the regulation of the proximal XOR promoter, it is of

interest that NF-Y A-subunit is up-regulated during the spontaneous enterocyte-like

differentiation of human Caco-2 colon carcinoma cells (Bevilacqua et al. 2002). Based on

this, we studied XOR expression during Caco-2 cell differentiation and, particularly, the role

of NF-Y in the process. A consensus GATA binding element, TGATAA, was identified in the

human XOR promoter on the region from –67 to –62. Except for the last nucleotide, the same

sequence can be identified in the rat promoter. Therefore, the interaction of GATA-4 and

GATA-6, both implicated in intestinal gene regulation, with XOR promoter was also

examined. The main results of the study are summarized in Table 3.

Expression of XOR during Caco-2 cell differentiation

The spontaneous differentiation of Caco-2 cells during a 15-day culture was assessed by

measuring maltase activity, which is a marker of enterocytic differentiation (Van Beers et al.

1995). There was no detectable XOR activity in conventionally subcultured subconfluent

Results

56

Caco-2 cells. However, after six days of culture measurable XOR activity appeared and was

quadrupled by day 15. A parallel induction of maltase activity was evident. Consistent with

XOR activity, a band corresponding to XOR protein became visible on day 6 and intensified

thereafter. XOR mRNA was faintly detectable already before measurable enzyme activity had

appeared and a more than fourfold increase in XOR mRNA levels ensued during the

differentiation process. -actin could not be used to control for the RNA content of the

samples, since its levels decreased during Caco-2 cell differentiation, and therefore 18S

ribosomal RNA was used to normalize the RNA content of the samples. The data suggest

transcriptional induction of XOR activity, even though a mechanism attributable to mRNA

stabilization can not be excluded.

Interaction of NF-Y with the XOR promoter is induced during Caco-2 cell differentiation

Protein binding to oligo1, carrying the NF-Y binding site, was apparent in nuclear extracts

isolated from undifferentiated Caco-2 cells on day 3. However, the band corresponding to

NF-Y binding to oligo1 was clearly intensified by day 6. Even though detectable on day 3, the

levels of NF-YA protein were elevated by day 6 and remained unchanged until day 15.

Binding of GATA-4 to the XOR promoter fragment XOR6E was suggested by EMSAs

performed with nuclear extracts derived from mouse Sertoli cells expressing follicle

stimulating hormone receptor (MSCFSHR-1) (Eskola et al. 1998) and showing high

endogenous expression of GATA-4 protein (Ketola et al. 1999), and from 293T cells

overexpressing GATA-4. Furthermore, supershift experiments with nuclear extracts derived

from undifferentiated Caco-2 cells indicated binding of GATA-6 protein, in addition to

GATA-4, to oligo2 carrying a GATA consensus element and spanning the nucleotides –92 to

–60 of the XOR promoter. (III, Fig. 4A to C). Neither binding to oligo2 nor the levels of

GATA-4 and GATA-6 proteins, changed during Caco-2 cell differentiation. We concluded

that NF-Y may be relevant in the transcriptional induction of XOR expression during

enterocyte-like differentiation of Caco-2 cells.

Results

57

Table 3. Summary of the results on maltase activity, XOR, NF-Y, and GATAs during Caco-

2 cell differentiation.

Day 3 6 8 12 15

Maltase activity + + + +++ +++

XOR activity + + ++ +++

XOR protein /+ + ++ +++

XOR mRNA + ++ ++ +++ +++

NF-Y binding toXOR promoter + +++ +++ ND +++

NF-YA protein + +++ +++ +++ +++

GATA-4/-6 protein + + + + +Undetectable ( ), barely detectable ( /+), detectable (+), between lowest and highest detected

(++), highest detected (+++), not determined (ND).

Hypoxic induction of human XOR activity is mediated by a

post-transcriptional mechanism (IV)

The proposed role of XOR in the pathogenesis of I-R injury together with the contradictory

results on the level of XOR induction in hypoxia, and, on the other hand, data suggesting

inhibition of XOR activity during hyperoxia, prompted us to study the regulation of the

human XOR under varying oxygen levels.

Hypoxia induces XOR activity, but does not alter XOR transcription or protein levels

Compared to BEAS-2B cells cultured in 21% O2 (normoxia), the XOR activity in cells

grown in 0.5% or 3% O2 (hypoxia) for up to 48 h was six- or eightfold, respectively.

However, XOR protein and mRNA levels as well as the proportion of XO of the total activity

remained unchanged. Furthermore, XOR promoter constructs were not activated by hypoxia,

whereas HRE-luc, used as a positive control for hypoxia, clearly responded. In agreement

Results

58

with this finding, cobalt, known to activate HIF-1, did not have an effect on XOR activity.

Substrate induction by hypoxanthine was excluded.

Hyperoxia inactivates XOR by a post-translational mechanism

XOR activity was markedly reduced in cells cultured in hypoxia followed by normoxia,

when compared to cells grown continuously in hypoxia (IV, Fig. 4). An analogous decrease in

XOR activity, but not in XOR protein, was noted when cells initially cultured in normoxia

were exposed to 95% O2 representing hyperoxia. Based on these findings, we assumed that

ROS may have a role in the inactivation of XOR. Indeed, the addition of iron into the culture

medium reduced the activity, but not the protein levels, of XOR overexpressed in 293T cells

(Figure 6, unpublished data), which is consistent with a role for the hydroxyl radical in XOR

inactivation. Function of the FAD center of XOR, assessed by measuring NADH oxidation,

remained intact in hyperoxia, but the DCPIP reducing activity was diminished by H2O2

rendering the Mo center as the likely target of inactivation.

Figure 6. Effect of iron on the activity and protein levels of XOR overexpressed in 293T cells.

The cells were transiently transfected with pcDNA3-XOR, and 17 h after the transfection either

normal medium (control) or medium with ferric ammonium citrate (FAC) was applied for a further 24

h of incubation. XOR (filled bars) and XO (open bars) activities were determined. To detect XOR

protein, western blot analysis with anti-XOR antibodies was performed. On each lane, 2.5 g of

protein were loaded, and -tubulin antibody was used as a loading control. Data are means of three

samples, and two of the samples were analysed on the gel. A representative experiment is shown.

Results

59

XOR expression is transcriptionally induced by iron, but XOR activity is

decreased by iron chelation at the post-translational level (V)

We studied the effects of intracellular iron levels on endogenous XOR in mouse NIH-3T3

and human BEAS-2B cells, on human XOR promoter constructs, and on XOR overexpressed

in cell culture. The influence of hydroxyl radical scavengers on the regulation of XOR by iron

was examined.

Increased intracellular iron induces XOR activity at the transcriptional level

A more than tenfold increase in intracellular iron resulted in duplication of the XOR

activity in NIH-3T3 cells after 24 h incubation (V, Fig. 1A and 1B). Parallel increases in

XOR protein and mRNA levels were apparent and the proportion of XO of total XOR activity

remained essentially unaltered. A less pronounced induction of activity was detected in

BEAS-2B cells. Inhibition of protein synthesis by cycloheximide inhibited the induction of

XOR activity, but not the elevation of XOR mRNA levels in NIH-3T3 cells incubated with

iron. Actinomycin D, used to inhibit RNA synthesis, abolished the induction of XOR activity

as well as the elevation of XOR mRNA levels. The potential role of the hydroxyl radical in

the signaling pathway leading to the induction of XOR transcription was evaluated by testing

the effects of hydroxyl radical scavengers on XOR activity. No effect on basal or iron induced

XOR activity was found (V, Table 2). Human XOR promoter constructs were not activated by

iron in transiently transfected 293T cells.

Iron chelation inactivates XOR by a post-translational mechanism

Data on the effects of iron chelator DFO on XOR are summarized in Table 4. DFO

decreased endogenous XOR activity in NIH-3T3 and BEAS-2B cells as well as the activity of

overexpressed XOR in 293T cells. The protein levels of XOR, however, remained unchanged.

The results indicate a post-translational inactivation of XOR activity by iron chelation.

Results

60

Table 4. Effects of 100 M DFO on XOR activity, protein and mRNA levels.

XOR Activity Protein mRNA

endogenous, NIH-3T3

endogenous, BEAS-2B ND ND

overexpressed, 293T ND indicates decrease, no change, increase, not determined (ND).

Discussion

61

DISCUSSION

Chromosomal localization of the human XOR gene

Our data on the chromosomal localization of the human XOR gene is in line with the other

reports defining the locus of the gene, which has been further confirmed at 2p23.1 by the

sequence data obtained in the human genome project. From an evolutionary point of view, it

is of interest that the human AO gene with remarkable sequence and structure homology has

been localized to chromosome 2q32.3 2q33.1 (Wright et al. 1995, International Human

Genome Sequencing Consortium 2001). Based on the localization in close proximity to each

other in human, the sequence homology, the conservation of exon-intron boundaries, and the

presence of XOR but the absence of AO-like activity in prokaryotes, XOR is likely to be the

precursor of a gene family, which has evolved through gene duplication and includes XOR,

mammalian AO, and the three recently identified mouse homologues of AO, named aldehyde

oxidase homologs (Terao et al. 1998; Terao et al. 2001; Garattini et al. 2003). Similarly, in the

same genetic region with human AO three pseudo-genes with homology to the AO gene have

been identified (Garattini et al. 2003).

In addition to the general interest of uncovering the physical location of a gene, and

possibly revealing multiple genes coding for a protein, determining the chromosomal

localization may allow conclusions concerning the regulation of gene expression related to the

state of chromatin. In yeast a correlation between the expression patterns of genes and

location on the same chromosome was demonstrated. Furthermore, functionally related genes

mapped to adjacent chromosomal regions, and it was speculated that common upstream

regulatory sequences may be shared by neighboring genes (Cohen et al. 2000). In agreement

with this, chromosomal clustering of highly expressed genes of specific human tissue samples

has been detected (Caron et al. 2001). Clustering of genes on the basis of tissue-specific

expression was also shown in C. elegans. This regulatory feature provided by the higher-order

organization of the genome was suggested to be based on the state of the corresponding

chromatin regions, facilitating the access of the transcriptional machinery or the utilization of

a common enhancer element (Roy et al. 2002). However, at present it is not known, whether

the XOR gene belongs to a physically linked cluster of genes expressed in a co-ordinate way

Discussion

62

under specific circumstances. Analysis of mRNA microarrays and functional studies based on

the sequence data of the human genome may provide novel insights into the relationship of

the chromosomal localization of the XOR gene and its regulation.

Regulation of the human XOR promoter

Our results from the functional characterization of the human XOR promoter constructs are

essentially in line with the data provided by Xu et al. (Xu et al. 2000). However, according to

our data the promoter fragment corresponding to nucleotides from –142 to +42 relative to the

translational initiation site showed the highest promoter activity, whereas Xu et al. reported a

marked increment in promoter activity along with the addition of nucleotides up to –224. This

discrepancy may be explained by differences in the contents of regulatory proteins in the cells

used by Xu et al. (HepG2 and human umbilical vein endothelial cells) and us (293T) to study

the promoter function. Alternatively, the presence of the first exon in the 3’-ends of our

constructs may result in promoter activities different from those obtained in the other study, in

which the 3’-ends of the constructs reached down to the nucleotide –1. Based on the data on

the rat XOR promoter function, there are crucial regulatory elements in the first exon (Clark

et al. 1998a), which shows considerable sequence homology (35 out of 42 nucleotides) with

the first exon of the human gene (Figure 7), suggesting that its presence may be important for

the proper function of the XOR promoter in human as well. Species characteristics may

account for the differences detected in promoter function, when mouse NIH-3T3 cells were

transfected.

Our data derived from the studies on DNA protein interactions and experiments with the

XOR promoter constructs in transiently transfected cells indicate that NF-Y binds to the

inverted CCAAT-box of the human XOR promoter and has an important functional role in the

basal regulation of the promoter. Binding of NF-Y was later confirmed with nuclear proteins

extracted from Caco-2 colon carcinoma cells undergoing enterocytic differentiation. These

results suggest that NF-Y may be involved in the up-regulation of XOR expression during the

differentiation process. The NF-Y binding site in the human promoter partially overlaps with

the C/EBP binding site of the rat promoter, which, however, does not interact with the

Discussion

63

enhancer binding factor I resembling NF-Y (Chow et al. 1995). We were not able to

demonstrate the binding of C/EBP proteins to the proximal human XOR promoter. The

location of the NF-Y binding site at –60 from the first suggested transcriptional initiation site

is in line with the available data on NF-Y regulated promoters (Mantovani 1998) and further

supports the role of NF-Y in the physiological regulation of the human XOR promoter. We

have also demonstrated the binding of GATA-4 and GATA-6 on the human XOR promoter. It

is tempting to think that these factors, which have been implied in the liver-specific gene

regulation as well as in the regulation of intestinal genes (Bossard and Zaret 1998; Molkentin

2000), might contribute to the regulation of the XOR gene. However, the functional

importance of GATA-4 and GATA-6 on the regulation of the XOR promoter remains

undetermined.

Xu et al. (2000) provided evidence on the binding of TFIID to the TATA-like element on

the human promoter region from –105 to –99, on the binding of a yet unidentified repressor to

an E-box (from –258 to –228), and on the repressive effects of these elements on the promoter

activity. We have not studied the relevance of these regulatory elements to the function of our

human XOR promoter constructs. In the rat promoter, no TATA-box has been identified.

However, despite the relatively high conservation of the coding sequences of XOR in human,

rat and mouse, the 5’-flanking sequence of the human XOR gene considerably differs from

the corresponding sequences in rodents (Cazzaniga et al. 1994; Chow et al. 1994), suggesting

that divergent transcriptional regulation may account for the species-specific regulation of

XOR expression. Figure 7 shows the putative proximal regulatory elements of the human

XOR promoter aligned with the corresponding rat promoter sequence. Furthermore, the 5’-

flanking sequences of the human AO show no homology to any of the XOR promoters (Terao

et al. 1998), which is in line with the different tissue distribution of the two enzymes. Taking

into consideration other elements of potential transcriptional importance, there exist, to our

knowledge, no data on putative regulatory elements in the introns or in the 3’-regions of any

XOR gene.

The existing functional data on the proximal human XOR promoter indicate that the

promoter activity is relatively weak under basal cell culture conditions. Enhancer elements

residing far away from the proximal promoter or even regulatory elements of other genes may

Discussion

64

be required for full transcriptional activity. In line with this, neither we nor others have been

able to directly show definite induction of the human XOR promoter under any specific

stimuli, even though transcriptional regulation is likely to occur, for example, when

intracellular iron levels increase. On one hand, the lack of strong and rapid transcriptional

induction may be related to the physiological role of XOR as a housekeeping enzyme of

purine catabolism. On the other hand, as described in this thesis, the regulation of the human

XOR gene expression takes place at several levels, transcriptional regulation representing

only one of them.

Figure 7. Human XOR promoter sequence from –224 to +42 aligned with the rat promoter. The

5’-ends of the XOR promoter constructs XOR4, XOR5, and XOR6 are shown. Inverted CCAAT

boxes, TATA box, and GATA element of the human promoter and two C/EBP binding sites of the rat

promoter are indicated. Rat sequence identical to the human sequence is shown in uppercase letters.

The potential transcriptional initiation sites are noted with downward arrows. The translational

initiation codon and the first exon are depicted.

Discussion

65

XOR expression during small-intestinal enterocyte-like cell differentiation

We have demonstrated the appearance of measurable XOR activity in a human colon

carcinoma Caco-2 cell line during the spontaneous small-intestinal enterocyte-like

differentiation of the cells. Along with the activity, elevations of the XOR protein and mRNA

levels were detected and an increased binding of NF-Y to an XOR promoter fragment was

found. Consistent with the available data on the levels of NF-Y subunits during Caco-2 cell

(Bevilacqua et al. 2002) and myeloid (Sjin et al. 2002) differentiation, we observed an

increment in NF-YA protein levels, suggesting that it may account for the increased NF-Y

DNA binding. Furthermore, the stable transfection of Caco-2 cells with NF-YA has been

demonstrated to result in the expression of small-intestinal differentiation markers

(Bevilacqua et al. 2002). Alternatively, post-translational modifications and interactions with

other proteins have been proposed to account for the increased NF-Y DNA binding capacity

during cell differentiation (Currie 1998b; Sjin et al. 2002).

In animal models the expression of GATA-4, -5, and -6 has been shown to vary in relation

to the location on the small-intestinal villi and maturity of the epithelial cells (Gao et al.

1998). Therefore, differential expression of the factors during intestinal differentiation was

considered possible. However, even though GATA-4 and GATA-6 are able to bind to the

human XOR promoter, no alterations in their binding to the promoter or in their protein levels

during Caco-2 cell differentiation were detected, the latter finding being in line with a

previous study (Fitzgerald et al. 1998).

Interestingly, NF-Y has been shown to control several genes involved in cell cycle

progression (Yun et al. 1999; Hu and Maity 2000; Manni et al. 2001), to regulate

ribonucleotide reductase (Filatov and Thelander 1995), and to interact with the c-myc proto-

oncogene (Taira et al. 1999; Izumi et al. 2001). On this basis, a role for NF-Y in the

regulation of cell proliferation and differentiation appears feasible. A histone deacetylase

inhibitor that prevents cancer cell growth and promotes differentiation requires an NF-Y

binding site to be able to transcriptionally activate thioredoxin-binding protein-2, the mRNA

levels of which are reduced in human colon and breast cancer (Butler et al. 2002). Also, two

other histone deacetylase inhibitors induce, by a mechanism involving NF-Y, the

Discussion

66

transforming growth factor type II receptor and thereby reverse the malignant phenotype

related to this gene in breast cancer cell lines (Park et al. 2002). The associations of NF-Y

mediated regulation with cell differentiation and malignant transformation are noteworthy

considering the regulation of human XOR by NF-Y and the appearance of XOR activity in

colon carcinoma cells undergoing enterocytic differentiation.

We have not been able to demonstrate measurable XOR activity in any human cancer cell

lines (unpublished data) and the same has been noted by others (Terada et al. 1997). Against

this background, the regulation of XOR by intracellular iron may be considered from another

perspective related to cell growth and differentiation. Iron is required for proliferation and has

been associated with the regulation of several proteins central for cell cycle progression, as

well as with genes involved in apoptosis such as p53, the proto-oncogenes N-myc and c-myc,

and with protein kinase C (Le and Richardson 2002; Templeton and Liu 2003). Furthermore,

iron is essential for the function of ribonucleotide reductase and iron chelation has been

shown to lead to the formation of a nonfunctional apoprotein (Cooper et al. 1996), which may

be analogous to the inactivation of XOR by iron chelation discussed in more detail below. To

our knowledge, the direct regulation of NF-Y by iron has not been studied despite the obvious

associations of NF-Y with processes regulated by iron.

Regulation of XOR by oxygen and iron

Our study on the regulation of XOR by oxygen suggests that the hypoxic induction of

XOR activity represents a post-translational reactivation of the enzyme and, on the contrary,

in ambient 21% O2 or in hyperoxia (95% O2) XOR is inactivated by oxygen metabolites. The

existing data on inactivation of XOR activity in hyperoxia and by ROS (Terada et al. 1988;

Terada et al. 1991; Hassoun et al. 1995) are in line with our results. However, enzyme

inactivation in 21% O2 was not detected in another study (Poss et al. 1996). Furthermore, two

studies reported a decrease of XOR mRNA along with the reduction of enzyme activity in

hyperoxia (Hassoun et al. 1994; Lanzillo et al. 1996), and conflicting data on the elevation of

XOR mRNA in hypoxia has also been provided (Hassoun et al. 1994; Lanzillo et al. 1996;

Poss et al. 1996; Terada et al. 1997). These discrepancies may be explained by the utilization

Discussion

67

of cells with varying growth rates from different sources and with differences in the capacity

to respond to elevated oxygen levels. Still another mechanism of hypoxic activation of XOR,

enhanced phosphorylation of the XOR protein, has been suggested to account for the

induction of XOR after 4 h incubation in 3% O2 (Kayyali et al. 2001). Based on our results,

increased phosphorylation can not be excluded, even though we did not notice XOR

activation by 4 h of hypoxic exposure.

We have confirmed the partial inactivation of purified XOR by H2O2 and, based on the

parallel inhibition of DCPIP reduction, which depends on an intact reduced Mo center,

provide evidence for the involvement of this redox active center in XOR inactivation.

Interestingly, the inhibition of XOR by NO results from a reaction with the critical sulfur of

the Mo center and subsequent formation of the desulfo-enzyme (Ichimori et al. 1999), which

may also happen upon inactivation of XOR by oxygen. The existence of a Mo cofactor

sulfurase, and the abscence of XOR activity in individuals lacking this enzyme, suggest a

physiologically relevant interconversion between the sulfo- and desulfo-XOR (Ichida et al.

2001), which may be the case in XOR inactivation by oxygen as well. Therefore, it will be of

interest to find out factors regulating the sulfurase itself. In order to further dissect the

mechanisms of oxygen inactivation, we showed that the FAD center of XOR remained intact

in cells exposed to hyperoxia. Iron-sulfur clusters are also potentially sensitive to oxygen

(Beinert et al. 1997), but since we did not study the function of the iron-sulfur clusters of

XOR separately, their role in XOR inactivation by oxygen cannot be ruled out.

Our data on the regulation of endogenous XOR by increased intracellular iron levels

indicate transcriptional activation of XOR. However, the addition of iron to 293T cells

decreased the activity of the overexpressed XOR by 30% without any change in the

immunoreactive protein levels. It is possible that iron induces oxidative stress and thereby,

together with ROS generated by XOR itself, inactivates the protein, as suggested by our data

on oxygen inactivation of XOR. Nevertheless, we suggest that under the control of the natural

regulatory elements of the endogenous XOR gene the transcriptional induction of XOR by

increased intracellular iron is the overriding mechanism. Consistent with this interpretation,

hydroxyl radical scavengers did not have an effect on endogenous XOR activity either in

Discussion

68

normoxia or in the presence of iron, which also suggests that signal transduction through ROS

may not be crucial for the transcriptional activation of XOR by iron.

A recently published study demonstrated an increase in XOR activity and protein levels in

cell culture after iron exposure and in the lung of rats exposed to iron (Ghio et al. 2002).

Consistently, iron chelation by DFO decreased XOR activity. In contrast to our results, the

rise in XOR activity in cell culture was detected already after 2 h of exposure to iron, and the

RNA synthesis inhibitor actinomycin D did not inhibit the induction of XOR activity by iron.

More precise mechanisms of XOR induction or inactivation were not determined. The authors

suggested that the rationale for the induction of XOR activity by iron may be the consequent

increased production of urate, which is an antioxidant and chelates iron, thereby providing

protection from iron-induced tissue injury (Ames et al. 1981, Davies et al. 1986). However,

considering this interpretation, the differences in the purine metabolism and physiological

urate levels between human and other mammals should be taken into account.

The lack of induction of XOR activity by cobalt and DFO is in line with the results

obtained from experiments on hypoxic regulation of XOR, suggesting that XOR is not an

HIF-1 responsive gene. The requirement of de novo RNA, but not protein, synthesis for the

induction of XOR activity by iron suggests that increased mRNA stability, mediated for

instance by IRPs, is an unlikely mechanism for the elevation of XOR mRNA levels.

However, a post-transcriptional mechanism of XOR mRNA stabilization by yet unidentified

proteins or regulatory RNAs can not be unequivocally excluded based on the application of

protein and RNA synthesis inhibitors. We did not study the more specific mechanisms of the

putative transcriptional induction of XOR, because our XOR promoter constructs did not

respond to changes in intracellular iron in 293T cells, suggesting that the appropriate

regulatory elements may not be present in our constructs. Alternatively, the effects of iron on

XOR activity may be cell-specific and some essential factors may be absent from the 293T

cells, which do not express endogenous XOR. Due to the low XOR promoter activity in NIH-

3T3 cells and inability to effectively transfect BEAS-2B cells, the transfection experiments

were conducted in 293T cells only. The inhibition of XOR activity by iron chelation, despite

an elevation in mRNA levels and unchanged protein levels, suggests that an apoprotein with

incomplete iron-sulfur clusters may be formed. This finding together with the data on the

Discussion

69

other post-translational modifications of XOR during hypoxia and hyperoxia, shows that the

levels of XOR mRNA and protein do not necessarily correlate with functional XOR protein

and underlines the importance of studying the regulation of gene expression at multiple levels.

Taken together, the existing data on XOR regulation by oxygen are compatible with the

following conclusions. First, hypoxia increases XOR activity and several mechanisms are

likely to be involved. Second, at a certain higher, but undefined, oxygen concentration

reduction in XOR activity ensues, and ROS contribute to the inactivation, which probably

results from the conversion of XOR into the inactive desulfo-form. Third, cell and tissue type

and their prevailing oxygen environment together with their antioxidative capacity and

different regulatory mechanisms presumably account for the degree of XOR activation in

biological systems. In addition, other factors, like increased intracellular iron, may modulate

XOR activity.

Pathophysiological aspects

In addition to the potential role of XOR in the development of I-R injury, XOR has been

ascribed a role in the pathogenesis of various diseases including coronary heart disease

(Spiekermann et al. 2003), heart failure (Cappola et al. 2001; Anker et al. 2003), hypertension

(Suzuki et al. 1998; Cai and Harrison 2000), inflammation and sepsis (Chinnaiyan et al.

2001), and liver diseases (Stirpe et al. 2002). In most of the human studies XOR activity has

not been measured, probably owing to the low or undetectable serum XOR activity in human,

but its significance has been evaluated indirectly by using XOR inhibitors. On the other hand,

based on the species-specific tissue distribution of XOR, the applicability of results obtained

in animal studies to humans has to be considered carefully.

Despite the physiological function of XOR in purine catabolism, it does not have a role in

the pathogenesis of gout, which is a common disorder of purine metabolism characterized by

elevated serum urate concentrations. However, inhibitors of XOR, allopurinol and oxypurinol,

which are structural analogues of hypoxanthine and xanthine, respectively, are commonly

used to treat gout (Becker 2001). No epidemiological data on increased mortality of patients

Discussion

70

with hereditary xanthinuria exists and approximately half of them appear completely healthy

(Raivio et al. 2001). In contrast, the XOR -/- mice die before the age of six weeks (Vorbach et

al. 2002), further emphasizing the species differences with respect to the physiology of XOR.

Specific findings of the XOR -/- mice have not been published. It will be interesting to find

out, whether XOR +/- mice, which are apparently healthy, except for lactation insufficiency,

present with any tissue pathology in the long term, and whether their ability to tolerate I-R

injury is different compared to normal mice.

Initially, the interest on XOR was mainly based on its putative contribution to I-R injury.

At least two critical requirements have to be fulfilled in order to ascribe XOR a role in the

pathogenesis of I-R injury. Firstly, XOR has to be present in the injured tissue and, secondly,

it has to be capable of producing harmful ROS. The latter prerequisite depends on the

prevalent form of XOR and on the relative amounts of the electron acceptors, NAD+ and O2.

In addition to XOR, there are several other potential endogenous and exogenous sources of

ROS, which may contribute to tissue injury involving increased ROS production and have to

be taken into account. These include the respiratory chain of mitochondria, nicotinamide

adenine dinucleotide phosphate (NADPH) oxidase in the plasma membranes of phagocytic

cells, irradiation, and some drugs (Kinnula et al. 1995). Conceivably, the regulation of XOR

activity by oxygen is important in the evaluation of the significance of XOR for ROS-

mediated tissue injury. Our data on inactivation of XOR in hyperoxia and reactivation in

hypoxia does not provide a definite answer to the potential contribution of this mechanism to

I-R injury. However, as total XOR activity is induced in hypoxia, a potential for increased

ROS production exists, assuming that eventually there is O2 available to be used as an

electron acceptor or that the XDH form donates electrons to O2 in the presence of reduced

amounts of NAD+. Whether the same applies to the subsequent reperfusion and reoxygenation

depends on the extent and direction to which XOR activity is influenced by oxygen

concentrations later, and therefore on the physiological levels of O2 in a specific tissue.

Furthermore, as the iron content of ischemic tissues increases, an induction of XOR activity in

prolonged hypoxia by an iron-related mechanism may follow along with the exacerbation of

tissue damage.

Discussion

71

In addition to I-R injury, the regulation of XOR activity by iron may be important in

chronic iron overload states exemplified by hereditary hemochromatosis. As a matter of fact,

increased body iron stores have been associated with an elevated risk for adverse

cardiovascular events (de Valk and Marx 1999). Interestingly, DFO, which chelates iron and

reduces XOR activity, has been shown to improve endothelial function in coronary artery

disease (Duffy et al. 2001). On the other hand, increased XOR activity was suggested to

contribute to endothelial dysfunction in coronary artery disease and to advance atherosclerosis

(Spiekermann et al. 2003). In line with this, XOR has been shown to bind to vascular

endothelium and inhibit NO-dependent vascular relaxation through the generation of O2

which further reacts with NO (Houston et al. 1999; Aslan et al. 2001). Cytokines have been

shown to induce XOR, and ROS produced by XOR have been suggested to contribute to

inflammatory response (Dupont et al. 1992; Falciani et al. 1992; Pfeffer et al. 1994). The

harmful effects of XOR on the endothelium may be worsened by the up-regulation of XOR

activity by cytokines, especially interferon- , which has been implicated in the atherosclerotic

process.

Our data on the appearance of XOR expression during Caco-2 colon carcinoma cell

differentiation suggests a link between XOR activity and cell differentiation. There are no

extensive data on XOR expression in human cancer, but disappearance of XOR protein was

detected in a small study on malignant breast tumors (Cook et al. 1997) and XOR activity was

clearly reduced in hepatocellular carcinoma when compared to normal liver tissue (Stirpe et

al. 2002).

Conclusions

72

CONCLUSIONS

In these studies, the chromosomal localization of the human XOR gene was determined,

and factors affecting the regulation of its expression under basal cell culture conditions,

during intestinal cell differentiation, by oxygen, and by iron were explored. The main

conclusions are:

1) The human XOR gene is localized on chromosome 2p22, which is in line with the

suggested evolutionary background of the XOR gene.

2) NF-Y has an important functional role in the transcriptional activation of the human XOR

gene. GATA-4 and GATA-6 also bind to the human XOR promoter, but their functional

significance to the transcriptional regulation of the XOR gene remains unproven.

3) Expression of XOR is induced at the transcriptional level during the enterocytic

differentiation of human colon carcinoma Caco-2 cells. A role for NF-Y is suggested, and

is in accordance with the putative association of XOR and cell differentiation.

4) Oxygen inactivates the XOR protein by interfering with the active site of the enzyme, and

the apparent induction of XOR by hypoxia is a reversal of this inactivation. Along with

the associations of varying oxygen levels and XOR activity, the transcriptional induction

of XOR activity by iron may contribute to the potentially detrimental effects of XOR in

I-R injury.

Acknowledgements

73

ACKNOWLEDGEMENTS

This study was carried out at the Research Laboratory of the Hospital for Children andAdolescents, University of Helsinki. I express my sincerest gratitude to Emeritus ProfessorJaakko Perheentupa, the former Head of the Hospital for Children and Adolescents, ProfessorMartti Siimes, the current Head of the Clinic for Children and Adolescents, University ofHelsinki, and Professor Erkki Savilahti, the Head of the Research Laboratory of the Hospitalfor Children and Adolescents, for providing excellent research facilities.

I am most grateful to my supervisors Professor Kari Raivio, the Chancellor of theUniversity of Helsinki, and Docent Risto Lapatto. I have been priviledged to work under theguidance of Kari Raivio, who has taught me the principles of scientific reasoning and writing.I want to thank him for trusting me and offering me independence in conducting my research.Still, he has always been ready to help me. His optimism and encouragement have been ofgreat value when facing difficulties. I admire Risto Lapatto’s enthusiastic attitude towardsscience in general, and I am most thankful for his support and interest in my research. Hisprofound knowledge on basic science and especially on protein chemistry has been of greatvalue during this project.

I want to warmly thank Docent Jorma Palvimo, whose vast knowledge on molecularbiology and expertise on the mechanisms of transcriptional regulation have beenindispensable during this work. As a member of my thesis committee, he has constructivelyevaluated the progression of my project and has always been ready to help. His advice andencouragement have been invaluable. I am grateful to Dr. Sirpa Leppä, the other member ofthe thesis committee, for her interest in my research project. I admire her energy as she, inaddition to her duties as a clinician and as a mother of three small children, manages to run aresearch laboratory.

Docent Pekka Kallio and Docent Ari Ristimäki are gratefully acknowledged for criticallyreviewing this thesis. Their analytical comments substantially improved the thesis, and I thinkour discussions have helped me prepare for the public examination.

I owe my warmest gratitude to Professor Markku Heikinheimo for collaboration, for hishelp with various problems, and for his interest in the scientific education of the students inour laboratory. Dr. Ritva Halila is warmly thanked for teaching me the basics of molecularbiology and for her friendship. The chromosomal localization of the XOR gene was “fished”in the research laboratory of Professor Aarno Palotie, under his and Dr. Maris Laan’s friendlyand most helpful guidance. I have also been priviledged to collaborate with Professor Olli-Pekka Kallioniemi.

I want to warmly thank the current and former members of our research group forfriendship and for the relaxed and supportive working atmosphere. Nina Linder has sharedwith me the joys and sorrows related to XOR, and to many other, also non-scientific, issues. Iam grateful to Mika Saksela for sharing his wide knowledge on XOR and for always beingready to help with problems of any kind. I have been lucky to have colleagues and friends likeTerhi Ahola, Henrikka Aito, Tiina Asikainen, Anna-Liisa Levonen, and Petra Pietarinen-Runtti. I owe special thanks to Anna-Liisa and Tiina, who have shared their experience and

Acknowledgements

74

helped me with many practical things and advice related to the preparation of the thesis. Terhiis especially thanked for professionally and patiently relieving the worries of a novice mother!

I am deeply grateful to Ritva Löfman and Sari Lindén for expert technical assistance andguidance. I wish to warmly thank Ritva for patiently teaching me the practical skills of thebasic methods in molecular biology. Sari’s help has been indispensable especially withHPLC, and her exact way of working has done the interpretation of data easier. It has alwaysbeen a pleasure to work with Ritva and Sari, and their support and encouragement have beeninvaluable during the years in the lab.

I wish to thank all the personnel of the Research Laboratory of the Hospital for Childrenand Adolescents for contributing to this work by generously offering their help. Thesupportive atmosphere in our laboratory has been of great importance especially during themoments of disappointment and frustration. My warmest thanks go to my dear colleagues MiaWesterholm-Ormio, Annaleena Anttila, Minna Eriksson, Krista Erkkilä, Sanne Kiiveri,Susanna Mannisto, Marjut Otala, Virve Pentikäinen, Laura Suomalainen, Ilkka Ketola, andJaakko Patrakka for their friendship. Kirsi Paukku and Pipsa Saharinen from the neighbouringlaboratory are thanked for helpful and amusing discussions.

Satu Mustjoki is warmly thanked for her friendship since the first day of the MedicalSchool. I greatly appreciate her views, and her encouragement has been irreplaceable. I amgrateful to all my friends for their sympathy, support, and for the enjoyable moments spenttogether.

My parents Paula and Jorma Rytkönen are sincerely thanked for everything. My mother’sendless optimism and my father’s faith in me have been invaluable during this, as well as anyother project in my life. I wish to express gratitude to my sister Liisi Klobut, my brother KariRytkönen, my sister-in-law Irmeli, and my brother-in-law Krzysztof for being there for me. Ihave always been welcome to their homes, where I have spent most joyful and relaxingmoments with my niece Anna and nephews Tuomas, Eelis, and Henrik. My parents-in-lawRitva and Kaj Martelin are acknowledged for their support and interest in my work.

Finally, my deepest thanks go to my husband Marcus for his love, continuous support, andpatience with me all these years. Our little son Eero helps me remember what really isimportant in life.

This work was financially supported by the Helsinki Biomedical Graduate School, theResearch Fund of the Helsinki University Central Hospital, the Foundation for PediatricResearch, University of Helsinki, and the Finnish Medical Foundation.

Helsinki, December 2003

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