REGULATION OF THE HUMAN XANTHINE OXIDOREDUCTASE GENE...
Transcript of REGULATION OF THE HUMAN XANTHINE OXIDOREDUCTASE GENE...
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
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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
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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
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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
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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
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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
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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
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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
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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
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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.
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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
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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.)
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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
Review of the Literature
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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).
Review of the Literature
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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
Review of the Literature
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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).
Review of the Literature
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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).
Review of the Literature
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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
Review of the Literature
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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
Review of the Literature
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.)
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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.
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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.
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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).
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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
Review of the Literature
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;
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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
Review of the Literature
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
Review of the Literature
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
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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).
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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).
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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,
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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
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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).
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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.
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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.
Review of the Literature
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
Review of the Literature
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
Review of the Literature
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
Review of the Literature
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
Materials and Methods
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
Materials and Methods
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
Materials and Methods
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).
Materials and Methods
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.
Materials and Methods
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
Materials and Methods
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
Materials and Methods
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
Materials and Methods
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.
Materials and Methods
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.
Results
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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