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A novel role for hypoxia-inducible factor-1alpha (HIF-1alpha) in the regulation of inflammatory chemokines and leukotriene expression in sickle cell disease

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A novel role for hypoxia-inducible factor-1alpha (HIF-1alpha) in the regulation of inflammatory chemokines and leukotriene expression in sickle cell disease

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A NOVEL ROLE FOR HYPOXIA- INDUCIBLE FACTOR -1α (HIF-1α) IN THE
REGULATION OF INFLAMMATORY CHEMOKINES AND LEUKOTRIENE
EXPRESSION IN SICKLE CELL DISEASE.

by

Caryn Suzanne Gonsalves

A Dissertation presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(BIOCHEMISTRY AND MOLECULAR BIOLOGY)

December 2009

Copyright 2009 Caryn Suzanne Gonsalves
ii

Acknowledgements
I am grateful to Dr. Kalra, my committee chair and mentor for giving me
the opportunity to work in his lab. Furthermore, my sincere thanks to Dr. Tahara,
for serving as a committee member and also for time spent answering my
numerous questions.
I also extend my appreciation to Dr. Robert Farley, Dr. Raymond Mosteller
and Dr. Robert Stellwagen, for their participation on my committee.
Further, I would like to thank Dr. Vikram Rajagopal, Dr. Shweta Shahi, Dr.
Kyoung-Soo Kim and Samantha Yeligar for their support and assistance.

iii

Table of Contents
Acknowledgements ii
List of Tables v
List of Figures vi
List of Abbreviations viii
Abstract xiii
Chapter I: Introduction 1
Pathophysiology of Sickle Cell Disease. 1
Hypoxia induced genes and role of HIF-1α in their
regulation. 3
Von Hippel Lindau (VHL) protein and PHD 2 regulate
HIF-1α expression under hypoxic conditions. 6
MicroRNAs and their role in gene regulation. 8

Chapter II: Hypoxia mediated up-regulation of 5-Lipoxygenase
activating protein (FLAP) requires HIF-1α. 11
Introduction 11
Materials and methods 16
Results 23
Discussion 44

Chapter III: Endothelin-1 mediated Macrophage Inflammatory
Protein-1β expression requires HIF-1α, in a
non-hypoxic manner. 49
Introduction 49
Materials and methods 52
Results 58
Discussion 72
Chapter IV: Sickle cell patients show elevated levels of urinary
leukotrienes. 77
Introduction 77
Materials and Methods 78
Results 79
Discussion 81

iv

Chapter V: Conclusions. 83
Bibliography 86

v

List of Tables
Table 1 Primers used in the study of hypoxia mediated FLAP
expression. 22

Table 2 Primers used in ET-1 mediated MIP-1β expression
studies. 57

vi

List of Figures
Figure 1 Pathophysiology of vaso-occlusions in sickle
cell disease. 2
Figure 2 A schematic representation of the HIF-1α subunits. 6
Figure 3 Biosynthesis of miRNA. 10
Figure 4 Overview of the pathways involved in the biosynthesis
of leukotrienes. 13
Figure 5 Hypoxia induced FLAP mRNA expression in t-HBEC
and HPMVEC. 26
Figure 6 siRNA p38, p47
phox
and p65 attenuate FLAP
expression under hypoxic conditions. 27

Figure 7 Hypoxia mediated FLAP mRNA expression involves
HIF-1α. 29
Figure 8 Hypoxia augments FLAP- Luc promoter via HIF-1α. 331
Figure 9 FLAP promoter activity under hypoxic conditions. 34
Figure 10 Hypoxia increases HIF-1α protein expression in
t-HBEC cells. 38
Figure 11 miR- 135a and miR -199a-5p regulate post-
transcriptional expression of FLAP. 431
Figure 12 Signaling pathways involved in hypoxia mediated
FLAP expression. 46
Figure 13 ET-1 augments expression of MIP-1β. 60
Figure 14 ET-1 induces MIP-1β release. 63
Figure 15 ET-1 mediated expression of MIP-1β requires
HIF-1α activity. 644
Figure 16 ET-1 augments MIP-1β promoter activity via HIF-1α. 676
Figure 17 ET-1 increases levels of HIF-1α protein and HIF-1α
binding to the MIP-1β promoter. 69
vii

Figure 18 miRNA-195 is down regulated in THP-1 cells in
response to ET-1. 721

Figure 19 Signaling pathways activated in ET-1 induced MIP-1β
expression. 75
Figure 20 Analysis of LTB4 levels in urine samples. 80

viii

List of Abbreviations
3' UTR 3' untranslated region
5-HPETE 5-hydroperoxy-6,8,11,14-(E,E,Z,Z)- eicosatetranoic acid
5-LO 5- lipoxygenase
AA arachidonic acid
ACS acute chest syndrome
AP-1 activator protein-1
ARNT aryl hydrocarbon receptor nuclear translocator
ATP adenosine triphosphate
C/EBP CCAAT box-enhancer-binding protein
CBP CREB- binding protein
CCR5 chemokine (C-C motif) receptor 5
CCR8 chemokine (C-C motif) receptor 8
ChIP chromatin immunoprecipitation assay
C-TAD C-terminal transactivation domain
Dn dominant negative
DPI diphenyleneiodonium chloride
ix

DTT dithiothreitol
ECE-1 endothelin-converting enzyme
EGTA ethylene glycol tetraacetic acid
EMSA electrophoretic mobility shift assay
Epo erythropoietin
ET-1 endothelin-1
ET
B
receptor endothelin b receptor
FLAP 5-Lipoxygenase activating protein
GAPDH glyceraldehyde 3-phosphate dehydrogenase
HbAS sickle cell trait
HbSS sickle hemoglobin
HIF hypoxia inducible factor
HLH helix-loop-helix
HPMVEC human pulmonary microvascular endothelial cells
HRE hypoxia response element
ICAM-1 inter-cellular adhesion molecule-1
IL-3 interleukin-3
x

IL-8 interleukin-8
JNK c-jun N-terminal kinase
LPS lipopolysaccharide
LTB
4
leukotriene B
4
LTC
4
leukotriene C
4
MAP kinase mitogen-activated protein kinase
MAPEG membrane-associated proteins in eicosanoid and
glutathione metabolism superfamily
MCP-1 monocyte chemotactic protein -1
MIP-1β macrophage inflammatory protein-1β
miRNA microRNA
mTOR mammalian target of rapamycin
NADPH nicotinamide adenine dinucleotide phosphate
NF-κB nuclear factor κB
NO nitric oxide
N-TAD N-terminal transactivation domain
ODD oxygen degradation domain
PAS domain PER-ARNT-SIM domain
xi

PHD prolyl hydroxylase
PHT pulmonary hypertension
PI3K phosphoinositide 3-kinase
PLA
2
phospholipase A
2
PlGF placenta growth factor
PMN polymorphonuclear neutrophils
PMSF phenylmethylsulphonyl fluoride
PTEN phosphatase and tensin homolog
qRT-PCR quantitative real time polymerase chain reaction
RANTES regulated on activation normal T cell expressed and
secreted
RBCs red blood cells
RISC RNA-induced silencing complex
RPA RNase Protection assay
SCD sickle cell disease
siRNA small interfering RNA
SPE solid phase extraction
SSRBCs sickle red blood cells
xii

t-HBEC transformed human brain endothelial cells
TNF-α tissue necrosis factor-α
VCAM-1 vascular cell adhesion molecule-1
VEGF vascular endothelial growth factor
VHL von hippel lindau protein
WBCs white blood cells

xiii

Abstract
In sickle cell disease (SCD), low oxygen tension results in sickle cell
hemoglobin polymerization, causing the sickling of red blood cells, with
subsequent vaso-occlusions (137). In the present study, we determined the
cellular signaling mechanism by which (i) endothelial cells are activated in
response to hypoxia to release leukotrienes and (ii) monocytes are activated by
ET-1 to form proinflammatory cytochemokines, and (iii) whether urinary
leukotriene levels in Tg SCD mice and SCD patients were higher compared to
corresponding control mice and healthy subjects. Our studies show that hypoxia
(1% O
2
) increased mRNA expression of FLAP, a key component of leukotriene
catalysis (107), in endothelial cells. The hypoxia- induced FLAP expression
required the activation of PI3 kinase, p38 MAP kinase, NAPDH oxidase, and HIF-
1α. FLAP promoter analysis demonstrated that mutation of the HRE regions and
NF-κB binding regions abrogated FLAP promoter activity, under hypoxia. We
also showed that miRNA-199a-5p and miR-135a regulated FLAP expression in
endothelial cells. Endothelin-1 (ET-1) levels are elevated in the plasma of SCD
patients. We show that treatment of THP-1, a monocytic cell line, with ET-1
induced the expression of a chemokine macrophage inflammatory protein -1β
(MIP-1β), a potent chemoattractant of natural killer cell and T-cells(133, 134).
ET-1 mediated MIP-1β expression required the activation of PI3 kinase, p38
MAP kinase, JNK kinase and HIF-1α. Mutations in the five HRE regions of the
MIP-1β proximal promoter mitigated MIP-1β promoter luciferase activity. These
xiv

studies, for the first time, establish a novel role of HIF-1α in the expression of
leukotrienes and chemokines mediated by hypoxia and ET-1, respectively. We
examined whether the data obtained in vitro was relevant in vivo in sickle cell
disease. Analysis of urine samples from sixty SCD patients showed ~2-fold
increase in LTB
4
and cysteinyl leukotrienes compared to 15 healthy normal
volunteers. These results show that SCD patients have higher levels of
leukotrienes and inflammation, which could be attenuated by using
pharmacological inhibitors of HIF-1α, providing new avenues for therapeutic
treatment of inflammation, asthma and vaso-occlusions in SCD patients.

1

Chapter I: Introduction
Pathophysiology of Sickle Cell Disease
Sickle cell anemia is an inherited hemoglobinopathy. The homozygous
form is characterized by the inheritance of a mutant β-globin chain of
hemoglobin, in which the glutamic acid residue at position 6 is substituted with
valine. Sickle hemoglobin is designated as HbSS (87). It is also the most
common variant, with severe clinical manifestations. The heterozygous form of
the disease, with inheritance of one mutant β-globin chain, results in the benign,
sickle cell trait (HbAS). Other variants, such as HbC and HbE, occur due to
inheritance of heterozygous states of sickle hemoglobin and its interaction with
other types of hemoglobin (87). When sickle erythrocytes (SSRBCs) pass
through capillaries, they adhere to the endothelium resulting in blood flow
obstruction. As a consequence, tissue hypoxemia develops leading to
polymerization of hemoglobin and altered shape of the red blood cells (RBCs)
(70). Sickle RBCs also display abnormal ion transport properties, which leads to
cellular dehydration and contributes to HbS polymerization (87).
A common complication of sickle cell disease is acute chest syndrome
(ACS). ACS is the second most common cause of morbidity and mortality in
sickle cell patients. ACS has been defined as a new infiltrate on a chest
radiogram associated with one or more symptoms such as fever, cough or new
onset hypoxia (68, 137). Some of the more common causes of ACS include
2

microbial infection, fat embolism arising from ischemic/necrotic bone marrow, or
vaso-occlusion (as reviewed (68), Figure 1).

Figure 1: Pathophysiology of vaso-occlusions in sickle cell disease.
The diagram illustrates the causes of vaso-occlusions in sickle cell disease. HbS polymerization
acts as a trigger for the development of vaso-occlusions. Localized tissue hypoxia can cause
sickling of RBCs. Sickle RBCs also exhibit impaired ion transport causing cellular dehydration,
contributing to HbS polymerization. The resulting hemolysis of sickle RBCs causes anemia and
indirectly reduces levels of NO, causing vaso-dilation and contributing to tissue hypoxia. Tissue
hypoxia can activate both the endothelium and leukocytes, which are also activated by decreased
NO levels, ischemia/reperfusion injury, and by placenta growth factor (PlGF).The activation of
the endothelium may occur as a direct result of tissue hypoxia and possibly due to release of
cytokines from the activated leukocytes. This leads to increased adherence of PMN and sickle
RBCs to the endothelium causing vaso-occlusions. Figure was adapted from (88).

Progression of pulmonary disease to ACS is most commonly attributed to
vaso-occlusions. Vaso- occlusions may be caused by the mechanical obstruction
of the blood vessels by rigid sickle RBCs. Adherence of the sickle RBCs (49),
platelets, polymorphonuclear neutrophils (PMN) and monocytes to the vascular
endothelium also play a role in vaso-occlusions (19) (Figure 1). The adherence of
3

WBCs to the vascular endothelium occurs as a result of the activation of the
endothelium (9, 53). As a result (131), the activated endothelium expresses
higher levels of adhesion molecules such as P-selectin (91), laminin, VCAM-1
(37) and ICAM-1. The endothelium can be activated by infection, tissue hypoxia,
proinflammatory molecules such as interleukin-1β and TNF-α, and vaso- active
molecules such as vascular endothelial growth factor (VEGF), endothelin-1 (ET-),
placenta growth factor (PlGF) and nitric oxide (NO) (9).
In sickle cell disease, localized tissue hypoxia arises due to increased
adherence of sickle RBCs. Hypoxia is common in venular beds with sluggish
blood flow and where deoxygenation of sickle RBCs can occur (128, 137). The
adhesion of leukocytes (PMN and monocytes) to the vascular endothelium and
its diapedesis through the blood vessel has been well documented and is thought
to be one of the factors in causing hypoxia in vivo (136, 151). However, relatively
less is known about the mechanism by which hypoxia activates the endothelium
to cause adhesion and diapedesis.
Hypoxia induced genes and role of HIF-1α in their regulation.
Adequate oxygen levels are necessary for various critical cellular
processes. Therefore, cellular and systemic oxygen concentrations are tightly
regulated via pathways that affect the expression and activity of a number of
cellular proteins.
4

Hypoxia arises when demand for oxygen exceeds supply. Hypoxia elicits
a cellular response designed to increase the amount of oxygen delivered to
tissue, while altering other cellular processes such as ATP production by
anaerobic glycolysis (13). Extensive studies have been conducted on the
oxygen dependent induction of genes, such as the genes encoding erythropoietin
(Epo), VEGF, and glycolytic enzymes (120, 140).
A key regulator in the oxygen dependent transcription of genes under
hypoxic conditions is the hypoxia inducible factor or HIF-1. Specifically, HIF-1
binds hypoxia-response elements or HRE regions, present in the promoter region
of genes such as erythropoietin (121, 122), vascular endothelial growth factor
(VEGF) (34) and interleukin-8 (IL-8) (74). The HIF complex is active only as a
heterodimeric protein, with two subunits, HIF-1α and HIF-1β (also known as aryl
hydrocarbon receptor nuclear translocator or ARNT). The HIF-1α subunit has
been shown to be regulated by oxygen tension, whereas the HIF-1β subunit is
constitutively active. Both subunits are members of the basic helix-loop-helix
(HLH)- containing PER-ARNT-SIM (PAS) domain of transcription factors (138)
(Figure 2). The HLH and part of the PAS domains mediate heterodimer formation
between HIF-1α and HIF-1β. The HLH domain is also necessary for binding to
the DNA (65). Transcriptional activation and interaction of HIF-1α with co-
activators is mediated via two transactivation domains, present in the C-terminal
region of HIF-1α. The transactivation domains are designated as the N- terminal
transactivation domains (N-TAD) and the C- terminal transactivation domains (C-
TAD) (Figure 2). Regulation of HIF-1α under normoxic conditions occur through
5

the oxygen-dependent degradation domain (ODD) which partly overlaps the N-
TAD (7). The ODD domain contains prolyl residues, which are hydroxylated
during normoxia. This facilitates the binding of the Von Hippel Lindau (VHL)
protein, a negative regulator of HIF-1α.
Other members of the hypoxia- inducible factor family include HIF-2α and
HIF-3α, which have more restricted expression patterns. HIF-2α (also called
endothelial PAS domain protein 1 or EPAS-1) is structurally and functionally
similar to HIF-1α (Figure 2). HIF-2α is also induced by hypoxia, dimerizes with
HIF-1β and mediates transcriptional activity (65, 143). HIF-2α is thought to be the
prevalent form in endothelial cells and fibroblasts, whereas HIF-1α has been
shown to be the dominant protein in epithelial cells (115). The normoxic
regulation of HIF-2α is similar to HIF-1α and involves binding of the VHL protein.
However, both factors regulate distinct groups of genes in vivo (55, 112).
HIF-3α is structurally similar to both HIF-1α and HIF-2α (Figure 2).
However, it lacks motifs required for transactivation, seen in the C terminus of
HIF-1α and HIF-2α. HIF-3α is also induced by hypoxic conditions, dimerizes to
HIF-1β and can bind to the HRE motifs. However, HIF-3α has been shown to
suppress HIF-mediated gene expression responses to hypoxia in the kidney and
may be involved in the negative regulation of transcriptional responses to
hypoxia (46, 89) (Figure 2).
6

Figure 2: A schematic representation of the HIF-1α subunits.
Boxes mark functional domains. The amino acid residues required for the regulation of HIF-1α
and HIF-2α are marked. bHLH, basic helix-loop-helix; PAS, Per/ARNT/SIM domains; ODD,
oxygen dependant degradation domains; N-TAD, N-terminal activation domain; C-TAD, C-
terminal activation domain; L-ZIP, leucine zipper. Figure was adapted from (7)

Von Hippel Lindau (VHL) protein and PHD 2 regulate HIF-1α expression
under hypoxic conditions.

mRNA for the HIF-1α and HIF-1β subunits are constitutively and
ubiquitously expressed under normal or hypoxic conditions(120). Regulation of
the HIF-1α subunit occurs at the protein level and is well defined under both
hypoxic and normoxic conditions.
Under normal oxygen tension, the HIF-1β subunit is constitutively
expressed. However, the HIF-1α subunit is highly unstable, with a half-life of 5
minutes (7). The rapid degradation of HIF-1α is mediated via hydroxylation of
prolyl and asparagines residues. HIF-1α undergoes hydroxylation at two prolyl
residues, 402 and 562, which are present within a conserved motif, L-X-X-L-A-P
(A, alanine; L; Leucine; P, proline; X, any amino acid) (52). Hydroxylation of
prolyl residues at 402 and 562 mediates the binding of the Von Hippel Lindau
7

(VHL) protein. VHL is the substrate recognition component of an E3 ligase
complex, which targets HIF-1α for proteasomal degradation (62, 93).
The hydroxylation of the prolyl residues of HIF-1α is catalyzed by the HIF-
1α prolyl hydroxylase (PHD) group of enzymes. In mammals, three prolyl
hydroxylase enzymes have been identified and are designated as PHD 1, 2 and
3 (reviewed in (81, 90). The PHD group of enzymes belongs to the 2-
oxyglutarate dependent dioxygenase superfamily. Both oxygen (as dioxygen O
2
)
(96) and 2- oxyglutarate are utilized as co-substrates for PHD enzyme activity.
Ferrous (Fe
2+
) is also required for enzyme activity (50).
As oxygen is required for the hydroxylation reaction (142), the HIF-1α
prolyl hydroxylases cannot function under hypoxic conditions. Therefore, VHL is
unable to target HIF-1α for proteasomal degradation. This allows for
accumulation of HIF-1α in the cell, under hypoxic conditions. The HIF-1α subunit
translocates to the nucleus, where it dimerizes with HIF-1β, and binds to HRE
motifs present in the promoters of target genes (62).
Oxygen tension also regulates the transcriptional activity of HIF-1α. HIF-
1α transactivation requires binding of the C-terminal transactivation domain of
HIF-1α to transcriptional factors CBP and p300 (4). The binding between HIF-1α
and CBP and P300, is regulated by the hydroxylation of an asparagine residue
(Asn 803) of HIF-1α (82). The hydroxylation reaction is mediated by an
asparaginyl hydroxylase or Factor inhibiting HIF-1 (FIH-1) (88, 95).
Hydroxylation of the asparagine residue prevents the binding of CBP and p300 to
8

HIF-1α, which in turn prevents HIF-1α from mediating transcriptional activation
(82).
MicroRNAs and their role in gene regulation.
MicroRNAs (miRNAs) are a class of newly discovered, 20-25 nucleotides,
non-coding RNAs (3). miRNAs are involved in gene regulation at the post
transcriptional level by either targeting mRNAs for degradation or translational
repression (3, 152). A vast majority of miRNAs are evolutionarily conserved
among various species (103). About 1% to 2% of the eukaryotic transcriptome is
represented by miRNAs (79).
A multi step process converts the longer transcripts of a miRNA gene to
the smaller miRNAs (Figure 3). miRNA genes are initially transcribed to primary
miRNA (pri-miRNAs), by RNA polymerase II (84). The pri-mi-RNA forms a hairpin
shaped stem-loop secondary structure and then enters a microprocessor
complex, composed of an RNAse III endonuclease, Drosha and a cofactor
DGCR8/Pasha (24, 41, 44, 83). First, DGCR8 recognizes the stem-loop
structures and binds to the pri-miRNAs. Drosha then cuts both strands of the
stem loop, asymmetrically at sites at the base of the stem, releasing a 60-70 nt
pre-miRNA (24, 41, 44, 83). Exportin 5 then transports the pre-miRNA to the
cytoplasm (12, 86, 150). In the cytoplasm, the pre-miRNA is processed further by
Dicer, a second RNAse III endonuclease (42, 59, 72). Dicer cuts a double
stranded miRNA: miRNA* duplex, with 5’ phosphate and a 3’ 2 nt overhang from
the end of the hairpin structure stem ,which is about 20-25 nucleotide (153). The
9

miRNA: miRNA* duplex is unwound by a helicase into two strands, the mature
miRNA and the antisense strand designated as miRNA*. The anti sense miRNA*
is degraded by an unknown nuclease. The mature miRNA is incorporated into a
ribonucleoprotein effector complex, known as the RNA-induced silencing
complex or RISC (43, 73, 118). Through the RISC complex, the miRNA can
induce translational repression or degradation of mRNA by binding to perfect or
nearly perfect complementary binding sites on the 3’ UTR of the target mRNA.
The complementarity between the miRNA and mRNA determines the mechanism
of gene regulation. Perfect binding between the miRNA and mRNA is thought to
bring about cleavage of the mRNA and its subsequent degradation (8, 114).
However, imperfect binding between the mRNA and miRNA is thought to bring
about translational repression. The number of binding sites between the miRNA
and the target mRNA determines the degree of repression (21). Studies suggest
that miRNAs hamper the movement of ribosomes along the mRNA, thus causing
translational repression (17). A third mechanism by which miRNAs can regulate
gene expression has been suggested. Recent studies suggest that miRNAs may
cause the rapid removal of the poly(A) tail of mRNAs, therefore accelerating
mRNA degradation (38, 146).
miRNAs have been shown to play a role in a number of cellular processes
such as cell differentiation, proliferation and normal development of tissues in
animals (16, 69, 97). miRNAs are also involved in mediating cellular responses
to various stimuli such as hypoxia (78). The loss of normal miRNA expression, as
seen in the case of miR15/16 or miR142, could lead to oncogenesis, which
10

suggests a role for these miRNAs as tumor suppressors. Over-expression of
certain miRNAs could also contribute to oncogenesis, suggesting that miRNAs
may serve as oncogenes (94).

Figure 3: Biosynthesis of miRNA.
Pre-miRNAs are cut by Drosha, releasing a pre-miRNA. The pre-miRNA is exported to the
cytoplasm by exportin 5. In the cytoplasm, Dicer cuts a double stranded duplex from the pre-
miRNA. The duplex is unwound and the sense strand is incorporated into a ribonuleoprotein
effector complex known as the RISC complex. Through the RISC complex, the miRNA can
induce translational repression or degradation of the target mRNA.

11

Chapter II: Hypoxia mediated up-regulation of 5-Lipoxygenase
activating protein (FLAP) requires HIF-1α.
Introduction
In sickle cell disease (SCD), low oxygen tension results in the
polymerization of sickle cell hemoglobin, causing the sickling of red blood cells,
with increased adherence of sickle RBCs, platelets, PMN and monocytes to the
vascular endothelium (49). This leads to the occlusion of blood vessels, clinically
characterized by recurrent episodes of painful crisis, which culminates in the
development of acute chest syndrome (ACS)(19). Adherence of blood cells to the
vascular endothelium occurs due to the activation of the vascular endothelium in
response to oxidative stress such as hypoxia or inflammatory cytokines (74). The
interaction between the activated endothelium and SSRBCs has been shown to
generate oxidative stress (47, 48, 130).
A consequence of hypoxia and oxidative stress is the formation of lipid
inflammatory mediators such as leukotriene B
4
(LTB
4
) and cysteinyl leukotrienes
(LTC
4
, D
4
and E
4
). Leukotrienes are potent mediators of inflammation involved in
asthma, allergic rhinitis and acute lung injury (123). Leukotriene B
4
(LTB
4
) is a
potent chemoattractant responsible for priming neutrophils for migration from the
bloodstream, into the extravascular space (127) and increases vascular
permeability (80). Leukotrienes C
4
,E
4
and D
4
interact with high affinity G protein
coupled receptors, causing contractile responses such as bronchoconstriction
12

(141) and changes in vascular permeability in the lungs (33). LTB
4
levels have
been shown to be higher in SCD patients, with higher levels seen in vaso-
occlusive crises and ACS (124).
Stimuli such as hypoxia, oxidative stress, microbes or allergens activate
the leukotriene biosynthesis pathway (Figure 4). Either the cytosolic or secretory
isoform of phospholipase A
2
(PLA
2
) initiates the pathway by catalyzing the
hydrolysis of arachidonic acid (AA) from membrane phospholipids (as reviewed
in (106)). Both 5- lipoxygenase (5-LO) and 5- lipoxygenase activating protein
(FLAP) are required for the synthesis of leukotrienes (26). 5-LO catalyses the
oxygenation of AA to 5(S)-hydroperoxy-6, 8, 11, 14-(E, Z, Z, Z)-eicosatetraenoic
acid or 5-HPETE. The second reaction catalyzed by 5- LO and FLAP converts 5-
HPETE to the epoxide 5(S), 6(S)-oxido-7, 9, 11, 15(E, E, Z, Z)-eicosatetraenoic
acid or leukotriene A
4
. Leukotriene A
4
is then metabolized to leukotriene B
4
, by
the enzyme, leukotriene A
4
hydrolase (116). Leukotriene A
4
can also be
metabolized to leukotriene C
4
by the enzymatic activity of leukotriene C
4

synthase (116). A membrane bound γ-glutamyl transferase transfers a glutamic
acid residue to convert leukotriene C
4
to leukotriene D
4
. Leukotriene D
4
is in turn
converted to leukotriene E
4
, by a specific membrane bound peptidase,
accompanied by the loss of a glycine residue (33).
5-LO is a member of the family of arachidonate lipoxygenases. Human 5-
LO is a 78 kDa protein and requires both ATP and Ca
2+
for its activity. 5-LO
differs from other members of the lipoxygenase family as it requires a
13

coactivator, FLAP for its activity. The gene for 5-LO has been characterized, and
is approximately 82 kb in length and has 14 exons divided by 13 introns (36).

Figure 4: Overview of the pathways involved in the biosynthesis of leukotrienes.
Arachidonic acid is released from cellular phospholipids by phospholipase A2 (PLA2), which is
then converted by 5-LO and FLAP to leukotrienes A4. Leukotriene A4 is transformed by the
activity of LTA4 hydrolase or LTC4 synthase into LTB4 and cysteinyl leukotrienes (LTC4, D4 and
E4). HPETE, 5(S)-hydroperoxy-6, 8, 11, 14-(E, Z, Z, Z)-eicosatetraenoic acid.

14

The NH
2
terminal of the protein, is coded by exons 1-7 with the carboxyl terminal
coded by exons 8-14 (36). The protein has four domains, an amino terminal β-
barrel structure and a carboxy terminal structure consisting predominantly of α-
helices. The carboxy terminal region is highly conserved across plants and
animals. The carboxy terminal region contains two clustered patterns of amino
acids, termed the lipoxygenase iron binding signatures which contain histidine
residues. The histidine residues hold a non-heme iron atom (107) and makes up
the catalytic domain of the protein (27, 92).E IN PRESS
FLAP and LTC
4
synthase belong to the membrane-associated proteins in
eicosanoid and glutathione metabolism superfamily (MAPEG) (63). Unlike other
family members, FLAP is not modulated by glutathione nor does it have
enzymatic activity (30). However FLAP has been shown to bind AA. The human
FLAP protein is an 18kDa membrane protein, with four transmembrane helices
(α1 to α4) that are connected by two elongated cytosolic loops (C1 and C2) and
one short luminal loop (30). The FLAP gene is 31 kb in length and has five exons
and four introns (71). FLAP expression is known to be induced by interleukin-1
(IL-3), dexamethasone and tissue necrosis factor-α (TNF-α). FLAP activity is
essential for leukotriene synthesis. Initial studies suggested that FLAP is
primarily expressed in cells of myeloid origin such as monocytes, macrophages,
eosinophils and basophil, however other studies have shown that FLAP is also
expressed in endothelial cells (20).
15

Studies have shown that TNF-α and lipopolysaccharide (LPS) induce the
expression of FLAP in THP-1 monocytic cells (113, 123). Also, transcription
factor nuclear factor-κB (NF-κB) and CCAAT box-enchancer-binding protein
(C/EBP) were required for LPS mediated FLAP expression in THP-1 cells (123).
We showed significantly higher levels of FLAP and 5-LO in the peripheral blood
monocytes of sickle cell patients, compared to normal individuals (105). Placenta
growth factor (PlGF) augmented the expression of leukotrienes from monocytes
and also upregulated 5-LO and FLAP mRNA expression. PlGF mediated FLAP
expression occurred via activation of HIF-1α, in a manner independent of hypoxia
(105).
In this study, we examined the effect of hypoxia on FLAP expression and
therefore leukotriene formation in human endothelial cells. We utilized
transformed human brain endothelial cell line (t-HBEC) for ease of culture and
transfection. Our studies show that hypoxia (1% O
2
) augments FLAP mRNA
expression. Hypoxia induced FLAP mRNA expression involved activation of
NADPH-oxidase, PI-3 kinase, MAP kinase, NF-κB and HIF-1α. In silico analysis
of the proximal promoter of FLAP revealed the presence of four HRE sites (-975
/+12 bp), which were essential for hypoxia mediated FLAP expression.
Additionally, NF-κB sites were also required for FLAP expression, under hypoxic
conditions. Since hypoxia induced the mRNA expression of FLAP gene, we
examined the potential miRNAs that may be involved in post-transcriptional
regulation of FLAP. Our studies for the first time, to the best of our knowledge,
16

show the role of miR-135a and miR-199a-5p in regulating hypoxia mediated
FLAP mRNA expression
Materials and methods
Cell Culture and reagents.
Transformed human brain endothelial cells (t-HBEC) were cultured in
RPMI-1640 supplemented with 1M L-glutamine, 1X MEM-vitamins, 1X non-
essential amino acids, 25 mg endothelial cell growth supplement, 10,000 units
sodium heparin, 1x penicillin-streptomycin, 0.5M HEPES, 1mM sodium pyruvate
and 10% heat inactivated FBS. Human pulmonary micro-vascular endothelial
cells (HPMVEC) were grown in EBM-1 media, supplemented with EBM-2 bullet
kits (Lonza, Cologne, Germany). Cells were kept in serum free media overnight
prior to stimulations. Cells were pre-treated with pharmacological inhibitors for 30
minutes before stimulation with hypoxia.
Inhibitors and antibodies
Diphenyleneiodonium chloride (DPI), LY294002, PD98059, SP600125,
and SB203580 were obtained from Tocris Bioscience (Ellisville, MO). R59949
(diacyl glycerol kinase inhibitor) and sulfasalazine were obtained from
Calbiochem, NJ. Antibodies for HIF-1α, β-actin and conjugated secondary
antibodies were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). All
other reagents, unless otherwise stated were obtained from Sigma-Aldrich (St.
Louis, MO).
17

Promoter and over-expression constructs
PTEN over expression and PI3K dominant negative plasmids were gifts
from Dr. Debbie Johnson (Keck School of Medicine, University of Southern
California, Los Angeles, CA). The -3368FLAP-pGL3 full length FLAP promoter
constructs and the deletion constructs (-965FLAP-pGL3, -371FLAP-pGL3 and -
134-FLAP-pGL3) were generously provided by Dr. Timothy Bigby (113)(VA
Hospital San Diego, La Jolla, CA). Control siRNA and siRNA for HIF-1α and
PHD2 were synthesized at the Microcore facility at the Keck School of Medicine,
USC. Control siRNA and siRNA for p38, p47
phox
and JNK2 were obtained from
Santa Cruz Biotechnology (Santa Cruz, CA).
mRNA extraction and analysis by qRT-PCR.
Endothelial cells (t-HBEC and HPMVEC) were incubated in 1% O
2
in
Forma tissue culture incubator (with oxygen sensor) for various time points and
total mRNA was extracted using TRIzol reagent (Invitrogen Life Technologies,
Carlsbad, CA). qRT-PCR was run using the iScript One-Step RT-PCR kit with
SYBR Green as per manufacturer’s instructions (Bio-Rad, Hercules, CA) on ABI
PRISM 7900 (Applied Biosystems, Foster City, CA). 40 cycles of amplification
was carried out following reverse transcription at 950Cx10s and 600Cx30s.
Relative quantification (RQ) values for mRNA expression were calculated as 2
-
ΔΔCt
by the comparative Ct method(108), where ΔΔCt = (Ct target gene of treated
sample – Ct GAPDH of treated sample) - (Ct target gene of control sample - Ct
GAPDH of control sample).
18

miRNA extraction and purification
Approximately 5 x 10
6
cells were kept in serum-free media for 8 hours and
treated with 1% O
2
for 8 hours. miRNAs were isolated and purified using the
miRVANA kit (Applied Biosystems/Ambion, Austin, TX), according to
manufacturer’s protocol. miRNA levels were detected using miRNA specific
primers (Applied Biosystems/Ambion, Austin, TX) by qRT-PCR. Ct values
obtained were normalized to Ct values for 5S. Relative quantification (RQ)
values for miRNA expression were calculated as 2
–ΔΔCt
by the comparative Ct
method (108), where ΔΔCt= (Ct target miRNA of samples- Ct 5S of treated
sample) - (Ct target gene of control samples-Ct 5S of control sample).
Preparation of protein extracts
Cytosolic and nuclear extracts were prepared as previously described
(25). Approximately 5 x 10
6
cells were washed and resuspended in 500 µl of cell
lysis buffer (10 mM HEPES (pH 7.9), 100 mM KCl, 1.5 mM MgCl
2
, 0.1 mM
EGTA, 0.5 mM DTT, 0.5 mM PMSF, 0.5% Nonidet P-40, and 1 µl/ml protease
inhibitor mixture) and left on ice for 30 minutes to swell. The lysate was vortexed
for 5- 10 seconds and centrifuged for 1 minute at 10,000g. The supernatant was
discarded and the pellet resuspended in 100µl of nuclear lysis buffer (10 mM
HEPES (pH 7.9), 1.5 mM MgCl
2
, 420 mM NaCl, 0.1 mM EGTA, 0.5mM DTT, 5%
glycerol, 0.5 mM PMSF, and 1 µl/ml protease inhibitor mixture). The lysate was
vortexed intermittently for 60 minutes. Lysates were centrifuged at 10,000 X g for
19

10 minutes. Supernatants were collected as nuclear extracts. Protein
concentrations were determined using the Bradford method (14).
Western Blot analysis
Nuclear extracts were used to determine HIF-1α protein levels. 25 µg of
protein were run on an SDS-PAGE gel. Membranes were blocked with 5% non-
fat milk. HIF-1α was probed using an antibody specific for HIF-1α (1:250).
Membranes were stripped and re-probed for β-actin (1:2500) levels to determine
equal loading. Protein extracts were detected using Immobilon western reagents
(Millipore Corporation, Billerica, MA).
Electrophoretic mobility shift assay (EMSA)
Single stranded complementary oligonucleotides were biotin labeled using
a Lightshift Chemiluminescent EMSA kit (104) (Pierce, Rockford, IL) and
annealed in equimolar ratios for 1hr at 37°C. The DNA binding reaction with 5 µg
of nuclear protein extract, 5% glycerol, 5mM MgCl
2
, 50ng/μl poly(dI·dC), 0.05%
NP-40 and 0.5ng biotinylated probe was incubated at room temperature for
20min. The specificity of DNA-protein interaction was demonstrated using 50-fold
excess of unlabeled probe. The samples were then subjected to non-denaturing
6% polyacrylamide gel electrophoresis in 0.5X TBE, transferred to a Hybond-N+
nylon membrane (Amersham Biosciences, Piscataway, NJ) followed by detection
with streptavidin-HRP/chemiluminescence.

20

Mutagenesis of FLAP promoter
HRE binding sites and NF-κB site mutants were generated using the Quik-
Change site directed mutagenesis kit (Stratagene, Cedar Creek, TX). The wild
type -965 FLAP luciferase construct was used as a template. Primers used are
shown in Table 1. Mutations were confirmed by DNA sequencing.
Transient Transfections
t-HBEC were transfected with various siRNA constructs (50 nM),1 µg of
FLAP luciferase reporter constructs and 1 µg of β-galactosidase constructs using
the protocol for Nucleofector T (Amaxa Biosystems, Cologne, Germany). RPMI-
1640 was used as the nucleofector solution. t-HBEC were transfected with 60
pmol of anti –miRNA inhibitor (Applied Biosystems/Ambion, Austin, TX) and 1 µg
of miRNA overexpression constructs (Genscript, Piscataway, NJ) using the
nucleofection protocol. For siRNAs, the sense and antisense oligonucleotides
were annealed at 95
o
C for 1 hour as previously described (45). Transfected cells
were kept in serum free media for 3 hours and then stimulated with hypoxia for
various time points. For luciferase assays, cells were harvested and analyzed for
luciferase activity (Promega, Madison, WI) using a luminometer (Berthold
Technologies; Lumat LB 9501), for the light emitted during the initial 10 s of the
reaction. β-galactosidase activity was assayed by colorimetric assay (Promega,
Madison, WI). The data are normalized for β-galactosidase activity and
expressed as relative luciferase unit. For mRNA analysis, cells were lysed in
TRIzol and mRNA isolated as described above.
21

Chromatin Immunoprecipitation (ChIP) assay
t-HBEC (10
7
cells) were kept overnight in serum-free RPMI-1640, followed
by treatment with hypoxia for the indicated time points. ChIP analysis was
performed utilizing HIF-1α antibody as previously described (74). Briefly, cells
were fixed with formaldehyde, lysed and chromatin was sheared by sonication (6
pulses at 15sec each, 40% potency). The lysate was centrifuged at 12,000 rpm
for 10 min at 4
o
C. The supernatants were pre-cleared for 2hr at 4
o
C with Protein
A-Sepharose beads (Sigma-Aldrich, St.Louis, MO). Precleared supernatants
were immunoprecipitated with HIF-1α antibody or control normal rabbit IgG
antibody at 4
o
C overnight. The immune complexes, with protein A beads were
collected and washed sequentially with low salt buffer, high salt buffer and TE
buffer. DNA cross-links were reversed at 65
o
C overnight, and DNA was extracted
by phenol/chloroform/isoamyl alcohol followed by ethanol precipitation.
Immunoprecipitated DNA was air- dried and re- suspended in 100 µl of nuclease
free water. DNA was subjected to PCR amplification for 30 cycles under the
following conditions; 95
o
C for 30s, 58
o
C for 60s and 72
o
C for 120s, using primers
listed in Table 1. The PCR products were subjected to agarose gel
electrophoresis followed by densitometric analysis. The values were normalized
to input DNA.
22

Table 1: Primers used in the study of hypoxia mediated FLAP expression.
Statistical Analysis
Control and hypoxia treated cells were compared by Student’s t-test. One-
way ANOVA followed by Turkey-Kramer test was used for multiple comparisons
using the Instat-2 software program (GraphPad, San Diego, CA). Values of
p<0.05 were considered statistically significant.

23

Results
Hypoxia augments 5- lipoxygenase activating protein (FLAP) expression in t-
HBEC and HPMVEC.
As shown in Figure 5A, a time dependent (2, 4 and 8 hours) increase in
FLAP mRNA expression was observed, when t-HBEC cells were treated with
hypoxia (1% O
2
). Treatment with hypoxia for 8 hours showed ~4-fold increase in
FLAP mRNA expression. However, 5-LO levels could not be detected at the
same time points, by RPA or qRT-PCR.
Hypoxia induced FLAP mRNA expression requires activation of PI3 kinase, MAP
kinase and p38 MAP kinase.
In order to determine the signaling pathway involved, various pharmacological
inhibitors were used. These pharmacological inhibitors have been shown to be
specific for kinases in cell signaling pathways. Pretreatment of t- HBEC with
LY294002 (15 µM), specific for phosphoinositide 3-kinase (PI3 kinase), followed
by exposure to hypoxia for 8 hours, inhibited expression of FLAP by 65% ±3%
(Figure 5B). Pharmacological inhibitors for mitogen-activated kinase or MAP
kinase (PD98059, 10µM) and p38 MAP kinase (SB203580, 1µM), inhibited FLAP
mRNA expression by 83+/- 5 % and 66 +/- 1%, respectively. An N-terminal Jun
kinase (JNK) inhibitor (SP600125) had no significant effect (Figure 5B). These
results suggest that the hypoxia induced cell signaling for the expression of FLAP
24

involved, PI3 kinase, MAP kinase and p38 MAP kinase, but not N-terminal Jun
kinase (JNK).
Furthermore, diphenyleneiodonium (DPI, 10µM), an inhibitor specific for
nicotinamide adenine dinucleotide phosphate (NADPH) oxidase (100, 129),
attenuated FLAP mRNA expression by 100% ± 2% (Figure 5C). As shown in
Figure 5C, sulfasalazine (2 µM), which inhibits NF-κB, attenuated hypoxia
mediated FLAP mRNA expression by 86% ± 5%. R59949 (30 µM), a diacyl
glycerol kinase inhibitor and putative activator of HIF-prolyl hydroxylases, also
inhibited hypoxia mediated FLAP mRNA expression by 104% ± 5% (Figure 5C).
These data indicate that FLAP mRNA expression, when induced by hypoxia,
involved PI3 kinase, MAP kinase, p38 MAP kinase, NAPDH oxidase and
transcription factors, NF-κB and HIF-1α.
Experiments were also performed using primary human pulmonary micro-
vascular endothelial cells (HPMVEC) to determine whether similar results were
obtained in primary cultures. Treatment of primary human pulmonary micro-
vascular endothelial cells (HPMVEC) with hypoxia for 8 hours increased by 4-fold
FLAP mRNA expression (Figure 5D). HPMVEC cells were pretreated with PI3
kinase inhibitor (LY294002) and R59949, a putative HIF-1α inhibitor. As seen in
Figure 5D, treatment with hypoxia caused a ~ 4 fold increase in FLAP mRNA
levels, which were attenuated by LY294002 (94% ± 5%) and R59949 (104% ±
6%). The results showed that hypoxia mediated expression of FLAP mRNA in t-
25

HBEC and HPMVEC utilized the same signaling pathways, and thus t-HBEC
could be used as a model system for further studies.
As the effect of pharmacological inhibitors may be non-specific, t-HBEC
were transfected with dominant negative PI3 kinase (Dn- Δp85) and an over
expression vector for PTEN. Both Dn PI3 kinase and PTEN over expression
completely attenuated FLAP mRNA expression, upon treatment with hypoxia
(Figure 6A). Similarly, transfection with short interfering RNA (siRNA) for p47
phox
, a subunit of NAPDH oxidase, completely attenuated FLAP mRNA, under
hypoxic conditions (Figure 6B). siRNA for p38 MAP kinase also completely
mitigated hypoxia induced FLAP mRNA expression (Figure 6C). However,
scrambled siRNA for p38 MAP kinase did not have an affect on FLAP
expression. Also, as shown in Figure 6C, siRNA for JNK-2 did not affect on FLAP
mRNA expression.

t-HBEC was also transfected with siRNA for p65, required for the activity
of NF-κB. siRNA for p65 also completely abrogated FLAP mRNA expression,
when treated with hypoxia (Figure 6D). Taken together, these results support the
involvement of PI3 kinase, MAP kinase, p38 MAP kinase, NADPH oxidase and
NF-κB, in hypoxia mediated FLAP expression. However, JNK kinase is not
activated in hypoxia mediated FLAP mRNA expression.

26

Figure 5: Hypoxia induced FLAP mRNA expression in t-HBEC and HPMVEC.
(A) t-HBEC cells were treated with 1% O
2
for indicated time points. (B) t-HBEC cells were
pretreated with LY294002 (15 µM), PD98059 (10µM), SB203580 (1µM), and SP600125 (100
nM). (C) t-HBEC were treated with sulfasalazine (2µM), DPI (10 µm) and R59949 (30 µm) for 30
minutes prior to treatment with 1% O
2
, for 8 hours. (D) Pretreatment of HPMVEC with LY294002
(15 µM) and R59949 (30 µM) for 30 minutes, was followed by treatment with 1% O
2
for 8 hours.
Total RNA was isolated for all experiments and subjected to quantitative qRT-PCR. Data are
expressed as means ± SEM of three independent experiments. Rel. - Relative. *** p< 0.001, **
p<0.01, *p< 0.05, ns, p>0.05.
HPMVEC
27

Figure 6: siRNA p38, p47
phox
and p65 attenuate FLAP expression under hypoxic
conditions.
(A) t-HBEC were transfected with 1 µg of Dn PI3 kinase and PTEN overexpression plasmids
and (B) 50nM of siRNA for p47 phox, (C) p38 MAP kinase and JNK-2 kinase , (D) siRNA for p65
, followed by treatment with 1% O
2
for 8 hours. Total RNA were isolated from samples and
subjected to qRT-PCR. Data were expressed as means ± SEM of three independent
experiments. Rel. - Relative. *** p< 0.001, ** p<0.01, *p<0.05, ns, p>0.05.

28

HIF-1α is activated in the hypoxia induced expression of FLAP.
R59949, a putative activator of HIF-prolyl hydroxylases, abrogated FLAP
mRNA expression, indicating a possible role for HIF-1α in the regulation of FLAP
expression under hypoxic conditions. Therefore, we transfected t-HBEC with
siRNA for HIF-1α, HIF-2α and corresponding scrambled HIF siRNA. As seen in
Figure 7A, siRNA for HIF-1α reduced hypoxia induced FLAP mRNA levels to
basal levels. However, scrambled, control siRNA (scRNA) and siRNA for HIF-2α
did not affect FLAP mRNA expression (Figure 7A).
Similarly, HPMVEC cells were also transfected with siRNA for HIF-1α. siRNA for
HIF-1α abrogated FLAP mRNA expression in HPMVEC under hypoxic conditions
(Figure 7B). Scrambled siRNA, however did not affect FLAP mRNA levels
(Figure 7B). Transfection with siRNA for prolyl hydroxylase2 (PHD2), which
causes the stabilization of HIF-1α protein, resulted in a ~7 –fold increase in FLAP
mRNA levels, under normoxic conditions. However, scrambled PHD 2 siRNA did
not affect FLAP mRNA expression (Figure 7C). These results suggest that
hypoxia mediated FLAP mRNA expression involved HIF-1α, but not HIF-2α.
Hypoxia mediated up-regulation of FLAP requires proximal HREs and the NF-κB
site.
Since siRNA for HIF-1α attenuated hypoxia mediated FLAP mRNA expression,
we determined if this occurred via the presence of hypoxia response elements

29

Figure 7: Hypoxia mediated FLAP mRNA expression involves HIF-1α.
(A and B) t-HBEC (A) and HPMVEC (B) were transfected with 50 nM of siRNA for HIF-1α and
HIF-2α. (C) t-HBEC was transfected with 50 nM siRNA for PHD2. Total RNA was subjected to
qRT-PCR. Data is expressed as mean ± SEM of three independent experiments. Rel.-Relative.
*** p< 0.001, ** p<0.01, *p<0.05, ns, p>0.05.

HPMVEC
30

(HREs). HREs are putative binding sites for HIF-1α, present in the promoter
regions of several genes (34, 74).
Analysis of the FLAP promoter (-3368/+12 bp) showed that hypoxia
increased the activity of the FLAP promoter by 3.5 fold, compared to the
promoter-less pGL3 vector (Figure 8A). The -965/+12bp region of the FLAP
promoter showed ~ 5-fold increased activity, higher than that of the full length
promoter activity. However, the -593/+12bp promoter region showed >90%
reduced activity. Similarly, the -371/+12bp FLAP promoter construct and -
131bp/+12bp construct, also showed > 90% reduced activity (Figure 8A). These
results indicated that the -965/+12bp promoter region was the minimal promoter
element essential for hypoxia induced FLAP transcription, and was utilized for
subsequent studies.
The minimal region of the FLAP promoter (-965/+12 bp) essential for
hypoxia mediated FLAP expression, showed the presence of four potential
putative consensus HRE sites (RCGTG)(105) at positions -170 to -167, -251 to -
248, -517 to -514 and -635 to -632 bp, as shown in the schematics of Figure 8B.
In addition, there is also present an NF-κB binding site at -43 to -34 bp
and two C/EBP consensus sites, located at -36 to -28bp and -25 to -12bp,
relative to transcriptional start site. Bigby and his coworkers (123) have
previously shown that LPS induced FLAP expression in THP-1 cells required
both NF-κB and C/EBP in its promoter. As shown in Figure 8C, mutations in
each of the four HRE sites (HRE-M1, -170 to -167 bp; HRE-M2, -251 to -248bp;
31

HRE-M3, -517 to -514bp; and HRE-M4, -635 to -632 bp) completely attenuated
hypoxia induced FLAP promoter activity, when compared to wild type promoter.
Mutations in the NF-κB binding site (located at -43 to -34 bp) also completely
attenuated FLAP promoter activity (Figure 8C). To determine, whether hypoxia
utilized the same HREs in primary endothelial cells, HPMVEC cells were
transfected with the minimal, -965/+12 bp FLAP promoter. As shown, hypoxia
induced a 3.5 fold increase in FLAP promoter activity, in HPMVEC (Figure 8D).
Mutations in the HRE sites completely abrogated the FLAP promoter activity,
under hypoxic conditions (Figure 8D). These results indicate that all four of the
HRE sites and the NF-κB site are required for FLAP promoter activity, under
hypoxic conditions in HPMVES as was seen in t-HBEC.

32

Figure 8, Continued.

33

Figure 8, Continued.

Figure 8: Hypoxia augments FLAP- Luc promoter via HIF-1α.
(A) Deletion analysis of FLAP promoter. t-HBEC was co-transfected with the indicated deletion
constructs and β-galactosidase plasmid, followed by treatment with 1% O
2
for 8 hours. The
luciferase activity was normalized to the promoter less pGL3 vector. (B) Schematics of the FLAP
promoter, depicting the putative binding sites for NF-κB, HIF-1α and C/EBP. (C and D) Hypoxia
induced FLAP promoter activity required four HRE sites and the NF-κB site in both t-HBEC (C)
and HPMVEC (D). Both, t-HBEC and HPMVEC were co-transfected with indicated constructs and
β-galactosidase plasmid, followed by treatment for 8 hours with 1% O
2
. Both luciferase and β-
galactosidase activity were measured as described in ‘Materials and Methods’. The luciferase
activity was normalized to that of the untreated -965 FLAP-luc construct. Data are expressed as
mean ± SEM of three independent experiments. *** p< 0.001, **p< 0.01, ns, p>0.05.

As shown in Figure 9A, hypoxia induced FLAP promoter activity was
reduced by LY294002 (96 ± 2%) and PD98059 (69 ± 3%). Rapamycin, an
inhibitor of mammalian target of rapamycin (mTOR) also inhibited FLAP promoter
34

activity by 73 ± 2%. Reductions in FLAP promoter activity were seen for inhibitors
SB203580 (77 ± 3%), sulfasalazine (54± 3%), DPI (73 ± 3%) and R59949 (69 ±
2%). However, SP600025 did not inhibit FLAP promoter activity.
Moreover, siRNA for HIF-1α attenuated FLAP promoter activity by more
than 90% (Figure 9B), under hypoxic conditions. Control, scrambled siRNA for
HIF-1α did not show any effect on FLAP promoter activity (Figure 9B). Since
pharmacological inhibitors inhibited FLAP promoter activity, these studies
support the role of PI3 kinase, NAPDH oxidase and p38 MAP kinase, and HIF-
1α, but not JNK kinase in hypoxia mediated FLAP expression.

35

Figure 9, Continued.

Figure 9: FLAP promoter activity under hypoxic conditions.
(A) t-HBEC cells were co-transfected with the minimal -965/+12 bp FLAP promoter and β-
galactosidase plasmid, followed by pretreatment with the indicated pharmacological inhibitors for
30 minutes and 1% O
2
for 8 hours. (B) t-HBEC cells were co-transfected with -965/+12bp FLAP-
Luc constructs and b-galactosidase plasmid and with siRNA for HIF-1α, followed by treatment
with hypoxia for 8 hours. Luciferase and β-galactosidase activity were measured as described in
‘Materials and Methods’. The luciferase activity was normalized to that of the untreated -965
FLAP-luc construct. Data are expressed as mean ± SEM of three independent experiments. ***
p< 0.001, **p< 0.01, ns, p>0.05.

Hypoxia increases HIF-1α protein expression
Studies have shown that hypoxia increases HIF-1α protein levels but not
HIF-1α mRNA levels (57). Hypoxia treated samples showed an increase in HIF-
1α protein levels in the nuclear extracts of treated t-HBEC cells (Figure 10A).
36

LY294002, PD98059 and SB203580 attenuated HIF-1α protein expression. In
addition, rapamycin, DPI and sulfasalazine also inhibited hypoxia mediated
increases in HIF-1α protein levels (Figure 10A). These results suggest that
hypoxia also mediates an increase in HIF-1α protein levels, which requires
activation of NADPH oxidase, PI3 kinase and p38 MAP kinase.
Hypoxia augments binding of HIF-1α to the FLAP promoter
Next, we determined whether hypoxia caused increased binding of the
HIF-1 transcription complex to DNA. Oligonucleotides flanking the first HRE at
position -170 to -167 bp were utilized as a probe for EMSA. As seen in Figure
10B, lane 1 and 2, increased binding of the HIF-1 transcription factor to DNA was
observed in nuclear extracts from cells treated with hypoxia, compared to
untreated cells. Furthermore, nuclear extracts from t-HBEC cells pre-treated with
LY294002 and R59949 showed reduced binding (Figure 10B, lanes 3 and 4).
50- fold excess of unlabeled probe competed out HIF-1α binding to DNA, which
proved the specificity of binding of HIF-1α (Figure 10B, lane 5). A mutant HRE
oligonucleotide (CGTG to AAAG) showed negligible binding of the HIF-1α
complex to DNA, when compared to HIF-1α binding with wild type HRE (Figure
10B, lane 6). These results were confirmed, utilizing a ChIP assay (Figure 10C),
where binding of HIF-1α protein to the FLAP promoter in native chromatin was
observed. Hypoxia treated samples showed ~5 fold increase in expected PCR
product of 319 bp (lane 2, Figure 10C). The product size corresponded to FLAP
promoter region, -310 to + 9 bp containing two HREs. Pretreatment of cells with
37

inhibitors LY294002, R59949 and sulfasalazine, attenuated by ~60% the PCR
product (lanes 4, 5 and 6, Figure 10C). Input DNA samples, which were not
immuno-precipitated with HIF-1α antibody, showed equal amount of amplification
(lower panel, Figure 10C). Immuno-precipitation of the chromatin samples with
control rabbit IgG did not show any amplification (middle panel, Figure 10C).
Taken together, these results indicate that hypoxia increases HIF-1α binding to
the FLAP promoter, both in vitro and in vivo, to upregulate the transcription of
FLAP.
Identification of miRNAs involved in hypoxia mediated FLAP-1 mRNA
expression.
Previous studies suggest a role for miRNA in the regulation of various genes (as
reviewed in (152)). Since hypoxia induced the mRNA expression of FLAP gene,
we examined the miRNAs that may be involved in stabilization and degradation
of FLAP. Previous studies have identified several miRNAs regulated by hypoxia
(78) that may play a role in the regulation of HIF-1α in cancer cells. Therefore we
determined the effect that hypoxia may have on the expression of hypoxia
regulated miRNAs i.e. miR-20a, miR-155, miR-106a, miR-135a, miR-17-3p, miR-
199a-5p and miR-203 in endothelial cells. As seen in Figure 11B, a decrease in
the expression of all the miRNAs was observed, when compared to expression in
untreated cells. Particularly, the expression of miR-135a mRNA was reduced by
90% relative to cells exposed to normoxia. The expression of miR-199a-5p
mRNA was reduced by 60%, under the same conditions.
38

Figure 10: Hypoxia increases HIF-1α protein expression in t-HBEC cells.
(A) t-HBEC cells were pretreated with the indicated inhibitors for 30 minutes prior to stimulation
with 1% O
2
for 6 hours. Rapamycin was used at a final concentration of 10 mM. 25 µg of nuclear
extracts were subjected to western blotting. Bands were detected using an antibody to HIF-1α.
The membrane was stripped and reprobed with antibody to β-actin, to normalize loading. Protein
bands were detected by chemiluminescence, corresponding to their molecular weights: HIF-1α
(120 kDa) and β-actin (42 kDa). Data are representative of 2 independent experiments. (B) t-
HBEC was pretreated with indicated inhibitors for 30 minutes, prior to treatment with 1% O
2
for 4
hours. Nuclear extracts were incubated with a biotin-labeled probe, corresponding to the region (-
179 to –159 bp) of the FLAP promoter. This region contains the proximal HRE located at -170 bp
to -167bp. 50 fold excess of cold probe was added in lane 5. A probe containing a mutation of
the HRE site (-170 to- 160 bp) was also used (lane 6). Data are representative of two
independent experiments. (C) t-HBEC cells were pretreated with inhibitors, followed by 1% O
2
for
4 hours. Chromatin was isolated and immunoprecipitated using an antibody to HIF-1α (upper
panel) or control rabbit IgG (middle panel). Primers used for amplification of products are listed in
Table 1. The lower panel is the amplification of input DNA before immunoprecipitation. Data are
representative of two independent experiments.

39

Figure 10, Continued

HIF- 1α
Free Probe
HIF- 1α
β-actin
40

Figure 10, Continued.

Similar results were seen for miRNA levels in hypoxia treated HPMVEC
cells (Figure 11C). We transfected t-HBEC cells with anti-miR oligonucleotides
for miR-135a and 199a-5p to determine if these miRNAs are influential in the
regulation of FLAP mRNA. As shown in Figure 11D, transfection of t-HBEC cells
with anti-miR-135a induced FLAP mRNA expression by 8-fold under hypoxic
conditions compared to hypoxia treated cells alone. However, anti-miR-199a-5p
was only 50% effective compared to anti-miR-135a in hypoxia induced FLAP
mRNA expression (Figure 11D, lane 4). HPMVEC cells were also transfected
with anti-miRNAs for miR-199a-5p and miR- 135 (Figure 11E). Transfection with
anti-miR for miR-135a induced FLAP mRNA expression by 5 –fold, when
compared to hypoxia treated cells, in HPMVEC. Anti- miR-199 also induced
FLAP mRNA expression by 3-fold, compared to hypoxia treated cells (Figure
11E).These studies indicated that miR-135a and miR-199a-5p could modulate
41

the intracellular levels of endogenous miRs to upregulate the expression of
FLAP. We, then, performed a converse experiment, to determine how
overexpression of miR-135a and miR-199a-5p would affect hypoxia induced
FLAP expression. Overexpression of miR-135a in HBEC abrogated FLAP mRNA
below the basal level, while miR-199a-5p reduced FLAP expression to the basal
level (Figure 11F). Taken together, these results indicate the role of miR-135a
and miR-199a-5p in regulating hypoxia mediated FLAP mRNA expression, in
both HBEC and HPMVEC.

42

Figure 11, Continued.

HPMVEC
43

Figure 11, Continued.

Figure 11: miR-135a and miR-199a-5p regulate post-transcriptional expression of FLAP.
(A) Putative binding sites for miR-135a and miR-199a-5p in the FLAP 3’UTR. (B and C) miRNA
expression in t-HBEC cells (B) and HPMVEC cells (C) treated with hypoxia. miRNAs were
purified as described in ‘Materials and Methods’. (D and E) t-HBEC cells (D) and HPMVEC cells
(E) were transfected with 60 pmol of anti miRNA inhibitor. (F) t-HBEC cells were transfected with
1 µg of overexpression plasmids for miRNA-135a and miRNA-199a-5p. Total mRNA was isolated
and subjected to qRT-PCR. Data are presented as means ± SEM and are representative of three
independent experiments. Rel.-Relative. ***p< 0.001, ** p<0.01, ns, p>0.05.
HPMVEC
44

Discussion
In sickle cell disease patients, LTB
4
levels have been observed 2-fold
higher when compared to healthy controls (124). Levels of leukotrienes were also
found to be elevated in SCD patients during episodes of vaso-occlussion and
acute chest syndrome (124). Recently, studies showed a 2-fold increase in
urinary cysteinyl leukotriene levels in children and adults with SCD, during a
painful crisis (31, 32, 64).
In the present study, we show that hypoxia increases FLAP mRNA expression
and subsequently, leukotriene expression. The levels of 5-lipoxygenase (5-LO)
levels were low in transformed human brain endothelial cells (t-HBEC) and did
not significantly increase in response to hypoxia. It has been shown that
endothelial cells do not express 5-LO (15). It has been suggested that adhesion
of cells such as PMN to the endothelium, may contribute to leukotriene formation,
by utilizing the LTA4 intermediate derived from PMN and leukocytes, in a
process termed as transcellular biosynthesis (15).
Next, we examined the cellular signaling pathway involved in FLAP
expression, under hypoxic conditions. Pharmacological inhibitors for PI3 kinase
(LY 294002), MAP kinase (PD98059), and p38 MAP kinase (SB203580)
attenuated FLAP mRNA expression (Figure 12). In addition, inhibitors for NADPH
oxidase (DPI), NF-κB (sulfasalazine) and a putative HIF-1α inhibitor (R59949)
also inhibited FLAP mRNA expression. However, an inhibitor for N-terminal Jun
kinase (SP600125) did not have any significant affect on hypoxia induced FLAP
45

mRNA expression, thus indicating that the JNK pathway was not involved in
hypoxia mediated FLAP expression (Figure 12). Since pharmacologic inhibitors
can be non-specific, we utilized the RNA interference (RNAi)-mediated gene
knockdown approach to delineate the signaling pathway. Transfection of t-HBEC
with siRNA for p47
phox
, a subunit of NADPH-oxidase, p38 MAP kinase, p65 (a
NF-κB component), inhibited expression of FLAP mRNA. Transfection of t-HBEC
with a dominant negative PI-3 kinase and a PTEN over-expression contruct,
attenuated hypoxia mediated FLAP expression (Figure 12). Our results indicated
that hypoxia-induced FLAP mRNA expression involved activation of NADPH-
oxidase, PI-3 kinase, p38 MAP kinase, MAP kinase, NF-κB and HIF-1α.
However, transfection with JNK siRNA did not affect hypoxia-mediated FLAP
expression, indicating that JNK pathway was not involved in hypoxia mediated
FLAP expression.
As HIF-1α and its family member, HIF-2α, share domain structure and
bind to identical motifs (RCGTG) but differ in their transactivation domains (98),
we examined the role both proteins played in FLAP expression. Transfection of
t-HBEC with siRNA for HIF-1α, but not HIF-2α, abrogated FLAP mRNA
expression. This suggests that both HIF-1α and HIF-2α may mediate expression
of different genes, as previously suggested (55, 98).

46

Figure 12: Signaling pathways involved in hypoxia mediated FLAP expression.
Our results show that hypoxia mediated FLAP expression requires the activation of PI3 kinase,
p38MAP kinase, NADPH oxidase, NF-κB and HIF-1α.

Transfection with PHD-2 siRNA, which stabilizes HIF-1α protein, augmented
FLAP expression under normoxia. This data supports the role of HIF-1α in
hypoxia mediated FLAP expression. Previously, our lab showed that PlGF, an
angiogenic factor, increased FLAP expression via HIF-1α, independent of
hypoxia (105). PlGF mediated FLAP expression involved two HRE sites, but not
the NF-κB site in the FLAP promoter (105). In the present study, we observed
that hypoxia- induced FLAP expression required four HRE sites along with the
NF-κB site in the proximal promoter of FLAP. This was demonstrated by mutation
of each of the HRE sites and the NF-κB site. The binding of HIF-1α to the HRE
sites in the FLAP promoter was further demonstrated by EMSA and ChIP. The
12
47

results obtained, indicated that hypoxia mediated FLAP expression involved HIF-
1α and NF-κB mediated signaling for FLAP expression. It has been shown that
LPS mediated FLAP expression in THP-1 cells required both NF-κB and C/EBP
binding to the promoter (123).
Most recently, miRNAs have been shown to regulate gene expression by
binding to specific sites in the 3’UTR of selected genes (28, 67, 110). miRNAs
have been shown to affect innate immune response (131), Toll-like receptor
expression (18) and erythropoiesis (139). Of the several signature miRNAs
regulated by hypoxia, which have been characterized and target HIF-1α (22, 79),
we have identified miR-199 and miR-135 as a target for hypoxia-induced FLAP
mRNA expression. In silico analysis of the 3’UTR of FLAP mRNA showed the
presence of complementary binding sites for miR-20, miR-155, miR-106a, miR-
135a, miR-17-3p, miR-199a-5p and miR203. Among these miRNAs, miR-199a-
5p and miR-135a were significantly attenuated by hypoxia. Transfection with anti-
miR-199a-5p and anti-miR-135a oligonucleotides induced FLAP expression
under hypoxia by 4- and 8-fold, respectively, compared to normoxia.
Overexpression of miR-135a-5p and miR-199a reduced FLAP mRNA levels
under hypoxic conditions to the basal level. These studies showed the regulatory
role of miR-135a and miR-199a-5p in hypoxia mediated FLAP mRNA expression.
Our studies show that hypoxia augments the expression of 5-lipoxygenase
activating protein via activation of HIF-1α and NF-κB. FLAP is also post-
translationally regulated via miRNA-199a-5p and 135a. Post –translational
48

repression by miR-135a and miR-199a-5p in human endothelial cells provides
another novel mechanism by which expression of FLAP is regulated. We suggest
that hypoxia mediated leukotriene formation in conjunction with previously
identified role of PlGF in LT formation (18), likely contributes to inflammation and
vaso-occlusion in SCD. These studies provide a new therapeutic approach,
based on HIF-1α target sites, to ameliorate inflammation and vasoocclusion in
sickle cell disease.

49

Chapter III: Endothelin-1 mediated Macrophage Inflammatory
Protein-1β expression requires HIF-1α, in a non-hypoxic manner.
Introduction
Sickle cell patients show increased incidences of hemolytic anemia,
frequent infections (145) and intermittent episodes of vaso-occlusive crises (29,
35, 109, 145). The most common cause of mortality in SCD is acute chest
syndrome (ACS). ACS is characterized by fever, cough and infiltrates in the lung
(68). Studies suggest that micro-occlusions may play a role in the development
of ACS (137). Repeated episodes of ACS contribute to the development of
pulmonary hypertension in SCD patients (5, 23, 75). Pulmonary hypertension
(PHT) develops with increasing age, in both adults and children with SCD, and is
a risk for early mortality (39). The vascular endothelium may contribute to vaso-
occlusive crises due to increased adherence of the sickle RBCs to the
endothelium (10, 49). Changes in vascular tone may also contribute to vaso-
occlusive crises. Vascular tone is maintained by endothelial cells via the
synthesis of vaso-dilators such as nitric oxide (NO) (102) and the expression of
vaso-constrictors such as endothelin-1 (ET-1) and eicosanoids (101). Studies
suggest that sickle erythrocytes could also influence vascular tone (99). SCD
patients show abnormal vascular tone (99). There is evidence that hemolysis-
associated PHT in SCD (39) patients results from impaired nitric oxide (NO)
availability that develops from quenching of NO by free heme (104). SCD
patients show elevated levels of ET-1, with higher levels seen during periods of
50

ACS (40). Hypoxic conditions due to tissue micro-occlusions and increased
hemolysis contribute to increased levels of ET-1 being released from endothelial
cells (77, 104).
ET-1 belongs to the endothelin family of peptides which includes
endothelin-2 and 3 (60). ET-1 is secreted primarily by vascular endothelial cells
and is one of the most potent vaso constrictors known (60, 61). Hypoxia (56, 77),
endotoxins, growth factors, and vasoactive hormones have been shown to
contribute to ET-1 generation and release (1). In addition, ET-1 production can
be inhibited by endothelium derived nitric oxide, heparin and nitro vasodilators
(1).
The ET-1 gene encodes a precursor molecule that undergoes post
translational cleavage. The precursor molecule is a 212- amino acid peptide
called the pre-proendothelin-1. Removal of a short secretory peptide generates
pro endothelin-1. Furin then cleaves pro endothelin-1, generating the biologically
inactive 38- amino acid precursor, referred to as big endothelin-1 (51, 148). The
mature, 21- amino acid endothelin-1 peptide, is obtained when big endothelin is
cleaved by one of several endothelin- converting enzymes (ECE), ECE-1(147).
Macrophage inflammatory protein-1beta (MIP-1β) or CCL4 belongs to the
C-C family of chemokines (85). MIP-1β is released by epithelial cells (149),
monocytes and T cells (144). Like other members of the C-C family of
chemokines, MIP-1β serves as potent chemoattractant for natural killer cells
(134), T-cells (132, 133), lymphocytes (117) and dendritic cells (125, 126). MIP-
51

1β serves as a ligand for both the chemokine C-C receptor-5 (CCR5) (2, 111)
and chemokine C-C receptor-8 (CCR8) receptors (11). MIP-1β and its
homologue, MIP-1α, together referred as MIP-1, have been shown to induce
localized inflammation, characterized by an influx of polymorphonuclear
neutrophils (PMN) and accumulation of mononuclear cells. This suggests a role
for MIP-1β in mediating the influx of PMN into the alveolar space, a common
occurrence in acute chest syndrome in SCD patients.
Previous studies have shown that the expression of chemokines
(interleukin-1 (IL-8), monocyte chemoattractant protein-1 (MCP-1) and MIP-1β)
were significantly higher in mononuclear cells from SCD patients (119).
Therefore, we hypothesize that increased levels of MIP-1β could play a role in
the transmigration of monocytes and leukocytes across the alveolar epithelium,
thereby contributing to lung injury and the symptoms seen in acute chest
syndrome. Higher levels of endothelin-1(ET-1) may contribute to monocyte
activation, seen in SCD patients, inducing expression of chemokines such as
MIP-1β, contributing to inflammation in sickle cell disease.
In this study we examine the effect of ET-1 on the expression of MIP-1β in
human monocytes. We utilized THP-1 monocytic cell line for ease of culturing
and transfection. Our studies show that ET-1 (250 nM) increases expression of
MIP-1β mRNA. ET-1 induced MIP-1β mRNA expression involved activation of
PI3 kinase, MAP kinase, NADPH oxidase, p38 MAP kinase, JNK kinase and HIF-
1α. Five HRE sites were found during in silico analysis of the proximal MIP-1β
52

promoter. As ET-1 induced expression of MIP-1β, we also examined the role
miRNAs play in the post- transcriptional regulation of MIP-1β. The present study
shows for the first time, to the best of our knowledge, the role miR-195 plays in
regulating ET-1 mediated MIP-1β expression.
Materials and methods
Cell Culture and reagents.
THP-1 cells were cultured in RPMI-1640, supplemented with sodium
pyruvate, L- glucose, penicillin-streptomycin (100 Units/ml) and 10% heat
inactivated FBS. Prior to stimulation with ET-1, cells were kept in serum free
media over night. Cells were pre-treated with pharmacological inhibitors for 30
minutes before stimulation with ET-1.
Reagents, inhibitors and antibodies
Diphenyleneiodonium chloride (DPI), LY294002, PD98059, SP600125,
and SB203580 were obtained from Tocris Bioscience (Ellisville, MO). ET-1, BQ-
610, BQ-788, R59949 (diacyl glycerol kinase inhibitor) and sulfasalazine were
obtained from Calbiochem (Gibbstown, NJ). Antibodies for HIF-1α, β-actin and
conjugated secondary antibodies were obtained from Santa Cruz Biotechnology
(Santa Cruz, CA). All other reagents, unless otherwise stated were obtained from
Sigma-Aldrich (St. Louis, MO). ET-1 (250 nM) was re-constituted in 5% acetic
acid.

53

Promoter and over-expression constructs.
PTEN over expression and PI3K dominant negative plasmids were gifts
from Dr. Debbie Johnson (USC). Control siRNA and siRNA for HIF-1α and HIF-
2α were synthesized at the Microcore facility at the Keck School of Medicine,
USC. MIP-1β promoter constructs were a gift from Dr. Marvin Reitz, Jr.,
Columbia University, New York (6).
mRNA extraction and analysis by qRT-PCR.
3 x 10
6
cells were kept in serum free media overnight. Cells were treated
with ET-1 (250 µM) and total mRNA was extracted using TRIzol reagent
(Invitrogen Life Technologies). qRT-PCR was run using the iScript One-Step
RT-PCR kit with SYBR Green as per manufacturer’s instructions (Bio-Rad,
Hercules, CA) on ABI PRISM 7900 (Applied Biosystems, Foster City, CA). 40
cycles of amplification was carried out following reverse transcription at 95
0
Cx10s
and 60
0
Cx30s, utilizing primers listed in Table 2. Relative quantification (RQ)
values for mRNA expression were calculated as 2
-ΔΔCt
by the comparative Ct
method, where ΔΔCt = (Ct target gene of SCD sample – Ct GAPDH of SCD
sample) - (Ct target gene of control sample - Ct GAPDH of control sample).
RNase Protection Assay
RPA was performed on total RNA using a custom probe H1 comprising of
tissue necrosis factor-α (TNF-α), interleukin-1β (IL-1β), regulated on activation
normal T-cell expressed and secreted (RANTES), MIP-1β, monocyte
54

chemotactic protein-1 (MCP-1), interleukin-1(IL-8) and the housekeeping genes
L-32 and glyceraldehyde 3- phosphate dehydrogenase (GAPDH) (BD
Biosciences). Briefly, 5 µg of total RNA was incubated with
32
P- labeled probes
for 12-18 hours at 56
o
C. The RNA duplexes were treated with RNase mixture
(BD Pharmingen), followed by extraction with phenol-chloroform. Protected RNA
duplexes were then run on a 6% denaturing polyacrylamide sequencing gel. The
gel was exposed to x-ray film for 8 hours.
Preparation of nuclear extracts
Cytosolic and nuclear extracts were prepared as previously described.
Approximately 10
7
cells were washed and resuspended in 500 µl of cell lysis
buffer (10 mM HEPES (pH 7.9), 100 mM KCl, 1.5 mM MgCl
2
, 0.1 mM EGTA, 0.5
mM DTT, 0.5 mM PMSF, 0.5% Nonidet P-40, and 1 μl/ml protease inhibitor
mixture) and left on ice for 30 minutes to swell. The lysate was vortexed for 5- 10
seconds and centrifuged for 1 minute at 10,000 X g. The supernatant was
discarded and the pellet resuspended in 100µl of nuclear lysis buffer (10 mM
HEPES (pH 7.9), 1.5 mM MgCl
2
, 420 mM NaCl, 0.1 mM EGTA, 0.5mM DTT, 5%
glycerol, 0.5 mM PMSF, and 1 μl/ml protease inhibitor mixture). The lysate was
vortexed intermittently for 60 minutes. Lysates were centrifuged at 10,000 X g for
10 minutes. Supernatants were collected as nuclear extracts. Protein amounts
were estimated using the Bradford method (14).

55

Western Blot analysis
25 µg of nuclear extracts were run on an SDS-PAGE gel. Membranes
were blocked with 5% non-fat milk and probed for HIF-1α protein levels.
Membranes were stripped and re-probed for β-actin levels to determine equal
loading.
Electrophoretic mobility shift assay (EMSA) for transcription factor HIF-1α binding
Complementary oligonucleotides were biotin labeled using a Lightshift
Chemiluminescent EMSA kit (Pierce, Rockford, IL) and annealed in equimolar
ratios for 1hr at 37°C. The DNA binding reaction with nuclear protein extract
(5μg), 5% glycerol, 5mM MgCl
2
, 50ng/μl poly(dI·dC), 0.05% NP-40 and 0.5ng
biotinylated probe was incubated at room temperature for 20 minutes. The
specificity of DNA-protein interaction was demonstrated using a 50-fold excess of
unlabeled probe. The samples were then subjected to non-denaturing 6%
polyacrylamide gel electrophoresis in 0.5 X TBE, transferred to a Hybond-N+
nylon membrane (Amersham Biosciences, Piscataway, NJ) followed by detection
with streptavidin-HRP/chemiluminescence.
Mutagenesis of MIP-1β promoter.
HRE, NF-κB and AP-1 binding site mutants were generated using the
Quik-Change site directed mutagenesis kit (Stratagene, Cedar Creek, TX). Wild
type -1065/+43 bp MIP-1β luciferase constructs were used as a template.
56

Mutations were confirmed by sequencing. Primers used for mutagenesis are
listed in Table 2.
Transient Transfections
THP-1 cells were transfected with various siRNA constructs (50 nM) using
the protocol for Nucleofector V (Amaxa Biosystems, Cologne, Germany). The
sense and antisense siRNA oligonucleotides were annealed at 95
o
C for 1 hour
as previously described (74). Transfected cells were kept in serum free media for
3 hours and then stimulated with ET-1. 2 µg of luciferase constructs were
transfected using the Nuclefector V protocol. 3 x 10
6
cells were used for
transfection. Cells were harvested and analyzed for luciferase activity (Promega,
Madison, WI) using a luminometer (Berthold Technologies; Lumat LB 9501), for
the light emitted during the initial 10 s of the reaction. β-galactosidase activity
was assayed by colorimetric assay (Promega, Madison, WI). The data were
normalized for β-galactosidase activity and expressed as relative luciferase units.
57

Table 2: Primers used in ET-1 mediated MIP-1β expression studies.
Quantification of MIP-1β.
3 x 10
6
cells were incubated in serum-free media, with or without ET-1 and
inhibitors. Supernatants were collected at indicated time points and stored at -
80
o
C. The amount of MIP-1β released was assayed using specific Duo-Set
ELISA kits (R & D systems) according to manufacturer’s protocol. Levels of MIP-
1β were normalized to total protein levels. Total protein levels were estimated
using the Bradford method (14).

58

Results
ET-1 augments MIP-1β expression in human THP-1 cells.
As shown in Figure 13A, treatment of THP-1 cells with ET-1, caused a
time-dependent, (15 minutes, 30 minutes, 1 hour and 2 hours) increase in the
mRNA expression of MIP-1β, TNF-α, and IL-1β. A modest increase in mRNA
levels was seen for RANTES. MIP-1β showed a four fold increase in mRNA at 30
minutes and 1 hour. We also examined the dose dependent (50 nM to 1000 nM)
effect of ET-1 on THP-1 cells. Optimal effect was observed at 250 nM (data not
shown).
ET-1 induced MIP-1β expression requires the activation of PI3 kinase, MAP
kinase, p38 MAP kinase and JNK kinase.
Treatment of THP-1 with pharmacological inhibitors of PI3 kinase (LY
294002,15 µM) , MAP kinase (PD98075, 10 µM), and p38 MAP kinase (SB
203580,1 µM)), followed by treatment with ET-1 inhibited mRNA expression of
MIP-1β by 88% ± 5%, 104% ± 7% and 102% ± 8%, respectively (Figure 13B).
Inhibitors for NADPH-oxidase (DPI, 10 µM) and an N- terminal Jun kinase
inhibitors (SP600125, 100 nM) inhibited MIP-1β mRNA expression by
approximately by 96% ± 2% and 118% ± 1, respectively. As shown in Figure
13C, sulfasalazine, an inhibitor for NF-κB, inhibited MIP-1β mRNA expression by
113% ± 7%. This suggests a role for NF-κB in the ET-1 mediated expression of
MIP-1β. Pretreatment of THP-1 cells with R59949, a diacyl glycerol kinase
59

inhibitor and putative activator of HIF- prolyl hydroxylases, also inhibited MIP-1β
mRNA expression by 109% ± 1% (Figure 13C). Ascorbate, an activator of
PHD2, also attenuated FLAP mRNA expression by 110% ± 11%. BQ788 (200
µM), an ET
B
receptor antagonist inhibited MIP-1β mRNA expression by 117% ±
7%, whereas BQ610 (200µM), an ET
A
receptor antagonist, did not show any
significant inhibition of MIP-1β expression (Figure 13C), indicating that the
signaling pathway involves the ET
B
receptor. However, we did not observe
similar increases in HIF-1α mRNA levels at the same time points (data not
shown).

60

Figure 13: ET-1 augments expression of MIP-1β.
(A) THP-1 cells were treated with ET-1 (250 nM) for the indicated time points. LPS (100 ng/ml)
was used as a positive control. Since we used 5% acetic acid as a solvent for ET-1, we treated
THP-1 cells with 0.005% acetic acid, as a negative control. 5 µg of isolated total RNA was used
for an RPA analysis. (B and C) THP-1 cells were pretreated with (B) LY294002 (15 µM),
PD98059 (10 µM), SB203580 (1µM), SP600125 (100 µM) and (C) with sulfasalazine (2 µm), DPI
(10 µm), ascorbate (25 µm), R59949 (30 µm), BQ610 (200 µM) and BQ788 (200 µm) for 30
minutes, followed by stimulation with ET-1 (250 µm) for 30 minutes. Total RNA was isolated and
subjected to qRT-PCR. Data are expressed as means ± SEM of three independent experiments.
Rel. - Relative. *** p<0.001, ** p< 0.01, ns, p>0.05

61

Figure 13, continued

62

Figure 13, Continued.

As mRNA expression of MIP-1β was regulated by PI-3 kinase and the ET
B

receptor, we determined if similar effects would be seen at the protein levels. As
shown in Figure 14, THP-1 cells, treated with ET-1 for 24 hours showed a 2-fold
increase in the release of MIP-1β protein, compared to that of untreated cells.
The PI-3 kinase inhibitor (LY294002, 10 µM) inhibited MIP-1β release by 134% ±
3%. R59949, a putative HIF-1α inhibitor, and an ET
B
receptor antagonist,
BQ788, inhibited MIP-1β release by 125% ± 3% and 125% ± 0.44%, respectively
63

(Figure 14). These data indicate that ET-1 mediated MIP-1β expression involved
ET
B
receptor and activation of PI3 kinase, MAP kinase, p38 MAP kinase and
JNK kinase. As R59949 inhibited MIP-1β expression, HIF-1α may be involved in
its expression.

Figure 14: ET-1 induces MIP-1β release.
THP-1 cells were pretreated with indicated inhibitors for 30 minutes, prior to treatment with ET-1
for 30 minutes. The supernatants were collected and assayed for MIP-1β release by ELISA.
Data are expressed as means ± SEM of three independent experiments. ** p< 0.01

ET-1 mediated up-regulation of MIP-1β requires activation of HIF-1α.
As R59949, a putative inhibitor of HIF-1α, inhibited MIP-1β mRNA
expression, we transfected THP-1 cells with siRNA specific for HIF-1α, HIF-2α
and a control scrambled siRNA (scRNA). ET-1 induced MIP-1β mRNA
expression was attenuated to basal levels by siRNA for HIF-1α (Figure 15).
However, both siRNA for HIF-2α and the control siRNA , did not show any affect
64

on MIP-1β mRNA expression (Figure 15). These results suggest that HIF-1α,
but not HIF-2α, is involved in the ET-1 mediated MIP-1β mRNA expression.

Figure 15: ET-1 mediated expression of MIP-1β requires HIF-1α activity.
THP-1 cells were transfected with siRNA (50 pmol) for HIF-1α and HIF-2α, followed by treatment
with ET-1 for 30 minutes. RNA obtained was used for a qRT-PCR analysis. Data are expressed
as means ± SEM of three independent experiments. *** p< 0.001, ns, p>0.05

ET-1 induced up-regulation of MIP-1β requires proximal HRE sites.
Since ET-1 mediated MIP-1β expression was attenuated by HIF-1α
siRNA, we determined if hypoxia response elements (HREs) played a role. HREs
are putative binding sites for HIF-1α, present in the promoter regions of several
genes (34, 74), including ET-1(104) and FLAP (105). ET- augmented full length
(-1065/ +43 bp) activity by 20 fold, compared to the promoter less pGL3 vector
(Figure 16A).
65

Analysis of serial deletions of the MIP-1β promoter showed that the -
1065/+43bp region of the MIP-1β promoter showed the maximum activity, while
the -485/+43bp promoter region showed reduced activity, when compared to the
-1065/+43 bp MIP-1β promoter construct. Similarly, the -264/+43bp FLAP
promoter construct and -192bp/+43bp construct showed >90% reduced activity
(Figure 16A). Therefore, the -1065/+43 bp region of the MIP-1β promoter, was
the minimal promoter element essential for ET-1 mediated MIP-1β expression
and was utilized for further studies. This region of the MIP-1β promoter showed
the presence of five potential putative consensus HRE sites (RGCAC) (56) at
positions -189 to -192 bp, -317 to -320 bp, -437 to -440 bp, -473 to -476 bp and -
703 to -706 bp (Figure16B). In addition, an NF-κB binding site is present at -77 to
-87 bp and an AP-1 and CRE- like binding site, located at -91 to -109 bp (6),
relative to transcriptional start site. Dr. Marvin Reitz and coworkers (6) have
shown that MIP-1β transcription in T cells, is regulated by the inducible cAMP
early repressor (ICER) and requires the activity of the AP-1/CRE-like binding site
(6). As shown in Figure 16C, mutation of the HRE sites at -189 to -192 bp,
(HRE-M1) inhibited MIP-1β promoter activity by 48% ± 0.86%. An HRE mutation
at -317 to -320 bp (HRE-M2) attenuated MIP-1β promoter activity by 53% ± 1%.
Mutations at HRE 3 (-437 to -476 bp, HRE-M3) mitigated MIP-1β promoter
activity by 51% ± 2% and HRE 4 (-473 to -476 bp, HRE-M4) attenuated by 47%
±6% the promoter activity compared to wild type (-1065 bp) promoter activity
(Fig. 16C). The HRE 5 mutant at -703 to -706 bp, (HRE-M5) resulted attenuation
of ET-1 promoter activity by 66% ±3% (Figure 16C). These results suggest that
66

all five HRE sites in MIP-1β promoter are essential for the ET-1 induced
expression of the MIP-1β gene.

67

Figure 16, Continued.

Figure 16: ET-1 augments MIP-1β promoter activity via HIF-1α.
(A) Deletion analysis of the MIP-1β promoter. THP-1 cells were co-transfected with the indicated
deletion construct (1 µg) and β-galactosidase plasmid (1µg). (B) Schematic diagram of the MIP-
1β promoter (-1065/ +43 bp), indicating the positions of the HIF-1α sites, NF-κB, AP-1 and CRE-
like binding sites. (C) ET-1 induced minimal MIP-1β promoter activity through the HRE sites. Data
are expressed as the means ±SEM of two independent experiments. ***p< 0.001, *p<0.05, ns,
p>0.05

ET-1 increases HIF-1α protein expression.
68

Recently, work from our lab has shown PlGF increased levels of HIF-1α,
but under non–hypoxic conditions (104). Thus, we determined whether ET-1
affected HIF-1α protein levels. As shown in Fig. 17A, ET-1 treatment of THP-1
cells showed an increase in HIF-1α protein levels in the nuclear extracts.
LY294002 attenuated HIF-1α protein expression (Figure 17A, lane 3). In addition,
R59949 also inhibited ET-1 mediated increase in HIF-1α protein levels (Figure
17A, lane 4). These results suggest that ET-1 mediates an increase in HIF-1α
protein levels, which requires activation of PI3 kinase and HIF-1α, and is
independent of hypoxia.
ET-1 augments binding of HIF-1α to the HREs in the MIP-1β promoter.
To determine if HIF-1α binds to the HRE sites in the MIP-1β promoter, an
oligonucleotide probe flanking the HRE closest to the transcription start site was
utilized for an EMSA. As shown in Fig.17B, nuclear extracts from ET-1 treated
THP-1 cells, showed increased HIF-1α binding, when compared to untreated
cells. Nuclear extracts from cells pretreated with LY294002 and R59949 showed
reduced binding. A 50-fold excess of cold probe competed out HIF-1α DNA
binding (Figure 17B). Moreover, an oligonucleotide with a mutation in the HRE
region of the MIP-1β promote, showed negligible binding of HIF-1α, when
compared to binding to wild type oligonucleotide, in the nuclear extracts from ET-
1 treated THP-1 cell (Figure 17B, lane 6). This data suggests that ET-1 increases
binding of HIF-1α to the MIP-1β promoter, therefore increasing MIP-1β gene
expression.
69

Figure 17: ET-1 increases levels of HIF-1α protein and HIF-1α binding to the MIP-1β
promoter.
(A) THP-1 cells were pretreated with inhibitors prior to stimulation for 30 minutes with ET-1.
Nuclear extracts (25 µg) were used for western blotting, with an antibody to HIF-1α. The
membranes were stripped and reprobed with an antibody for β-actin to normalize for any loading
differences. (B) THP-1 cells were treated with inhibitors for 30 minutes, followed by treatment
with ET-1 for 15 minutes. Nuclear extracts (5µg) were incubated with a biotin-labeled probe,
containing the proximal HRE site (-189 to -192) in the MIP-1β promoter. In lane 6, 50 fold excess
of unlabeled probe was added. A probe with a mutation in the HRE site (-189 to -192) was also
used (lane 5).

70

Identification of miRNAs up-regulated upon ET-1 treatment.
Analysis of the 3’UTR of MIP-1β using the Sanger miRNA database,
revealed putative binding sites for a number of miRNAs. As shown in Fig. 18A,
3’UTR of MIP-1β showed complimentary binding sites for miR-195 and miR-
223. Based on this information we examined the effect candidate miRNAs 20a,
194, 195 and 223 on MIP-1β expression (Figure 18A). THP-1 cells treated with
ET-1 for 30 minutes showed a significant down regulation in the expression of
miRNA-20, 194 and 195 (Figure 18B). However miR-223 was up regulated in
response to ET-1 treatment. This data suggests a possible role for miR-20, 194
and 195 in the regulation of MIP-1β expression. miR-223 was up regulated in
response to ET-1. As miR-195 was upregulated in response to hypoxia and also
possibly plays a role in tumor development, we focused our studies on this
miRNA. THP-1 cells transfected with anti-miR-195 increased MIP-1β expression
by ~4 –fold, when compared to ET-1 treated cells (Figure 18C). These results
suggest a role for miR-195 in the post-transcriptional regulation of MIP-1β, in
response to ET-I treatment.

71

72

Figure 18,Continued.

Figure 18: miRNA 195 is down regulated in THP-1 cells in response to ET-1.
(A) Putative binding sites for miR-195 and miR-223 in the 3’ UTR of MIP-1β. Sites were
determined using the Sanger miRNA database. (B) MiRNA 195 and 194, but not miRNA 223, are
down regulated in response to ET-1 treatment. THP-1 cells were treated with ET-1. MiRNA was
extracted as described in ‘Materials and Methods’ and subjected to qRT-PCR. Data are
expressed as means ± SEM of three independent experiments. *** p < 0.001, ns, p>0.05

Discussion
Pulmonary complications are an increasing cause of mortality in SCD
patients. Incidences of restrictive lung disease (75, 76, 135) and pulmonary
hypertension (PHT) (5, 23, 39) increase with age. The pulmonary vascular tone
is maintained by the actions of ET-1, a vaso-constrictor and nitric oxide, a vaso-
dilator. Increased hemolysis seen in SCD patients (39, 66) and transgenic SS
73

mice(54), can decrease the availability of NO, which may contribute to the
increased incidence of pulmonary hyper tension. ET-1 levels are elevated in
sickle cell patients, with highest levels seen during acute chest syndrome (40).
Previous studies from our lab have shown that monocytes from SCD patients are
in a highly activated state and also express higher levels of cytokines (IL-1β and
TNF-α) and chemokines (MCP-1 and MIP-1β )(119).
In this report we show that ET-1 induced MIP-1β expression in monocytes.
For these studies, we utilized a monocytic cell line, THP-1, for ease of culture
and transfection. THP-1 cells treated with ET-1 showed a 4- fold increase in
expression of MIP-1β. Next, we examined the ET-1 mediated cellular signaling
pathway for the expression of MIP-1β. We observed pharmacological inhibitors
of PI-3 kinase (LY294002), MAP kinase (PD98059), p38 MAP kinase
(SB203580) and JNK kinase (SP600125) attenuated ET-1 mediated MIP-1β
mRNA expression. Additionally, NF-κB inhibitor (sulfasalazine), NADPH oxidase
inhibitor (DPI) and R59949, a putative inhibitor of HIF-1α abrogated MIP-1β
mRNA expression. Ascorbate, which causes the increased degradation of HIF-
1α, also inhibited MIP-1β mRNA expression. As the effects of pharmacological
inhibitors are non-specific, we utilized various siRNAs to delineate the signaling
pathway as depicted in schematics of Figure 19. Our results utilizing
pharmacological inhibitors specific for kinases along with gene knockdown
approach utilizing siRNAs revealed that ET-1-induced MIP-1β mRNA expression
involved activation of NADPH-oxidase, PI-3 kinase, p38 MAP kinase, MAP
kinase, NF-κB and HIF-1α. Since HIF-1α and HIF-2α share homology in protein
74

and domain structure, and bind to identical HRE motifs, thus we examined which
of these HIFs were involved in MIP-1β expression in response to ET-1.
Moreover, HIF-1α and HIF-2α differ in their transactivation domain (98). siRNA
for HIF-1α, but not HIF-2α attenuated the expression of MIP-1β mRNA in
response to ET-1. These results suggest a role for HIF-1α, but not HIF-2α in ET-
1 mediated MIP-1β expression, under non-hypoxic conditions.
Studies have shown the involvement of AP-1 complex in LPS mediated
MIP-1β expression (6). In silico analysis of the MIP-1β promoter revealed the
presence of five HRE sites in the reverse orientation (RGCAC) on the sense
strand and an NF-κB binding site at -77 to -83bp from the transcriptional start
site. We observed that ET-1 mediated MIP-1β expression required all five HREs
sites and NF-κB site in its promoter.
Further studies are warranted to determine if the AP-1 complex is also
essential for ET-I mediated MIP-1β promoter activity. The role of HREs in the
proximal promoter of MIP-1β was demonstrated by EMSA, wherein nuclear
extracts from ET-1 treated cells specifically bound HIF-1α to the oligonucleotide
probe containing HRE consensus sequence of MIP-1β.
Furthermore, inhibitors of PI3 kinase and HIF-1α attenuated the binding of
HIF-1α to the oligonucleotide probe. Mutations in the HRE site also abrogated
HIF-1α protein binding to the oligonucleotide. These results indicated that ET-1
mediated MIP-1β expression involved HIF-1α mediated signaling.
75

Figure 19: Signaling pathways activated in ET-1 induced MIP-1β expression.
Our studies suggest that MIP-1β expression requires the activation of PI3 kinase, p38 MAP
kinase, JNK kinase, HIF-1α and NF-κB.

Recently microRNAs (miRNAs) have been shown to regulate gene
expression (152). Analysis of the 3’UTR for MIP-1β, by the Sanger miRNA
database, showed putative binding sites for several miRNAs including miR-223,
194, 195 and 20a. We, therefore, investigated the expression levels of miRNAs-
20, 195, 223 and 194 in response to ET-1. ET-1 treatment of THP-1 cells down
regulated the expression of miR- 20a, 194 and 195 but caused an increase in the
expression levels of miR-223. We therefore hypothesize that miR-20a, 194 and
195 may play a role in regulation of MIP-1β, but not miR-223. Transfection with
anti-miR-195 increased MIP-1β mRNA expression by ~4 fold, when compared to
76

cells treated with ET-1. Further experiments include investigating the role of
miRNAs by utilizing over expression plasmids for miR-195.
Overall, these studies show that ET-1 plays a role in the expression of
MIP-1β, via activation of HIF-1α. We suggest that increased levels of ET-1 seen
during ACS may be responsible for the up-regulation of MIP-1β. Previous studies
have shown that MIP-1β serves as potent chemoattractants for natural killer
cells (134), T-cells (132, 133), lymphocytes (117) and dendritic cells (125, 126).
Higher levels of MIP-1β may be responsible for the increased transmigration of
leukocytes across the vascular endothelial and into the alveolar space, which is
commonly seen during ACS (136, 151). These studies provide a new target for
therapy of sickle cell disease, which would help ameliorate the inflammation and
other symptoms of ACS seen in SCD patients, utilizing inhibitors that inhibit or
degrade HIF-1α.

77

Chapter IV: Sickle cell patients show elevated levels of urinary
leukotrienes.
Introduction
Leukotrienes play an important role in airway inflammation and airway
hyperreactivity, symptoms seen in patients with acute chest syndrome.
Additionally, leukotrienes play an important role in inflammatory diseases such as
inflammatory bowel disease, asthma, and rheumatoid arthritis. Of the
leukotrienes, leukotriene B
4
, (LTB
4
) is a potent chemoattractant responsible for
priming neutrophils for migration from the bloodstream, into the extravascular
space (127) and causing an increase in vascular permeability (80). Leukotrienes
C
4
, E
4
and D
4
are collectively termed cysteinyl leukotrienes . The cysteinyl
leukotrienes interact with high affinity G protein coupled receptors causing
contractile responses such as bronchoconstriction (141) and changes in vascular
permeability in the lungs (33).
Studies done to determine the levels of leukotrienes in SCD patients
showed higher levels of LTB
4
in the urine and plasma (124). Higher levels of
leukotrienes may contribute to the inflammation and ACS seen in SCD patients.
Therefore, we seek to determine if increased levels of leukotrienes in sickle cell
patients can serve as an indicator for the severity of acute chest syndrome.
Leukotrienes (LTB
4
, LTC
4
, LTD
4
and LTE
4
) levels were measured in urine
samples of 100 patients with β-thalessimia, which showed significant (2-3 fold)
78

increases in LTB
4
and LTE
4
, but not in LTC
4
by statistical analysis (data not
shown). These studies established the methodology for measuring leukotrienes.
The levels of leukotrienes, thus determined, will be used for determining the
degree of severity of acute chest syndrome and asthma in sickle cell patients.
We hypothesize that increased leukotriene levels will correlate with degree of
asthma and acute chest syndrome in SCD patients. Thus measurement of levels
of leukotrienes in urine or plasma can be used as a prognostic indicator for the
degree of severity of acute chest syndrome in sickle cell patients. Furthermore,
levels of LTs will provide an indicator for the effectiveness of treatment with
leukotriene antagonists, such as Zileuton, for children and adults with SCD.
Materials and Methods
Purification of leukotrienes from urine samples.
Urine samples were obtained from patients in accordance with IRB
regulations, from the University of Cincinnati Medical Center, Cincinnati, Ohio.
Samples were purified for Leukotriene B
4
and cysteinyl leukotrienes (E
4
, C
4
, and
D
4
) using solid phase extraction columns (SPE columns). Briefly, 2 ml of the
urine (for human samples) and 500 µl (for mouse urine samples) were used for
purification. Protein was precipitated using equal volumes of ethanol (LTB4) or
methanol (cysteinyl leukotrienes). The samples were spun at 5000 rpm for 10
minutes. Supernatants were acidified to pH 4 using dilute hydrochloric acid.
Supernatants were purified using solid phase extraction columns (SPE columns,
79

Cayman chemicals). Briefly, the columns were activated by 5 ml of methanol,
which was followed by a wash with 5 ml water. Samples were then introduced
and allowed to flow through the columns. The columns were then washed with 5
ml of water and hexane, respectively. The columns were allowed to dry after the
hexane wash. The columns were then eluted with either 5 ml of ethanol (LTB
4
) or
5 ml of methanol (cysteinyl leukotrienes). The elutants were evaporated to
dryness under a stream of nitrogen and reconstituted with either 500 µl of assay
buffer for human samples or 200 µl of assay buffer for mouse samples. The
samples were diluted and assayed utilizing an EIA plate, as described in the
manufacture’s protocol (Assay Designs).
Creatinine Estimations
Creatinine values were estimated using Jaffe’s reaction (58). Briefly, 500
µl of urine samples were mixed with 500 µl of 0.75 N of NaOH and 500 µl of
0.1% of picric acid, and incubated for 20 minutes at 30
o
C. Absorbance values for
each of the samples were read at 490 nm. The data was computed utilizing
standard curve for creatinine.
Results
Initial studies on leukotriene levels in urine samples were done using
samples from normal and sickle cell transgenic mice. The sickle cell transgenic
mice have the human sickle cell gene for β-globin. Samples were obtained from
4 normal and 4 Tg sickle mice. Leukotriene B
4
levels were significantly higher (~
80

1.4 fold) in Tg sickle mice compared to normal mice, as shown in Figure 20A.
However, cysteinyl leukotriene levels did not show a significant difference
between the normal and Tg sickle mice (data not shown).
For the analysis of leukotriene levels in SCD patients, sixty samples were
obtained from SCD patients, while fifteen normal patients contributed urine
samples. Analysis of human urine samples showed 2-fold higher levels of
leukotriene B
4
in sickle cell patients compared to normal patients (Figure 20B).
Moreover, cysteinyl leukotriene levels were also ~ 4 fold higher in sickle cell
patients compared to normal healthy individuals (Figure 20C).

20A
81

Figure 20: Analysis of LTB4 levels in urine samples.
(A and B) LTB4 levels were significantly higher in sickle mice (A) and SCD patients (B) compared
to corresponding controls. (C) Cysteinyl leukotriene (Cyst LTs) levels were significantly higher in
sickle cell disease patients when compared to normal controls. n=4 for control and sickle mice,
n=15 for normal patients and n=60 for SCD patients.

Discussion.
Previous studies by Setty et. al (124) have shown elevated LTB4 levels in
sickle cell patients. In this study, we examined LTB
4
and cysteinyl leukotriene
levels in sickle cell patients and SCD patients with acute chest syndrome. The
purpose of this study was to determine if urinary leukotriene levels can serve as
20B
20C
82

an indicator for the degree of severity of ACS. Our data shows increased levels
of LTB
4
and cysteinyl leukotrienes in SCD patients with ACS. This validates the
data obtained from studies done by other labs. The data obtained from this study
will be used to correlate degree of severity of acute chest syndrome in sickle cell
patients with levels of leukotrienes. Further studies will examine the levels of
urinary leukotrienes in SCD patients treated with drugs that inhibit leukotriene
formation, such as montelukast (Singulair) or Zileuton and the extent of acute
chest syndrome in sickle cell patients. Zileuton is an inhibitor of 5-Lipoxygenase,
whereas montelukast is a leukotriene receptor antagonist. Both drugs are at
present used for the treatment of asthma. If significant decreases in the levels of
urinary leukotrienes are observed, in patients using Zileuton or montelukast,
these drugs could be used as an effective drug for the control of ACS in sickle
cell patients

83

Chapter V: Conclusions
In sickle cell disease, low oxygen tension causes the polymerization of the
sickle hemoglobin, causing the sickling of red blood cells (136). Sickle red blood
cells, along with platelets, PMN and monocytes, exhibit increased adherence to
the vascular endothelium bringing about the occlusion of blood vessels (137).
Clinically, vaso-occlusive crises are characterized by recurrent episodes of
painful crises (128). Adherence of these cells to the vascular endothelium occurs
in response to the activation of the endothelium by oxidative stress, such as
hypoxia or inflammatory cytokines (10). Previous studies have shown that
hypoxia upregulated the expression of inflammatory cytokines in endothelial cells
(74). Moreover, studies have shown that leukotriene levels are augmented in
sickle cell disease patients, which further increase during acute chest syndrome
(124). However, relatively less is known how hypoxia contributes to increased
formation of leukotrienes.
In our first study, we determined the mechanisms involved in the
activation of endothelial cells and the subsequent release of leukotrienes.
Hypoxia (1% O
2
) increased the expression of 5- lipoxygenase activating protein
(FLAP), a key catalytic component, required for leukotriene biosynthesis.
Hypoxia- mediated expression of FLAP required the activation of PI3 kinase, p38
MAP kinase, NADPH oxidase, NF-κB and HIF-1α. Furthermore, siRNA for p38
MAP kinase, p65 (a subunit of NF-κB complex) and p47
phox
, a subunit of NADPH
oxidase, attenuated FLAP mRNA expression. In addition, siRNA for HIF-1α also
84

abrogated FLAP mRNA expression. Mutations of the HRE and NF-κB binding
sites in the proximal promoter of FLAP also inhibited FLAP promoter activity.
These results show, for the first time, the role of HIF-1α in hypoxia mediated
FLAP expression. We also investigated the role of miRNAs in regulation of FLAP
gene expression. Our studies, demonstrate for the first time, the role of miR-135a
and miR- 199a-5p in regulating FLAP expression.
In our second study, we investigated the role ET-1 played in contributing
to the proinflammatory state in SCD. Previous studies have shown increased ET-
1 levels in the plasma of SCD patients (40). It has been suggested that elevated
ET-1 levels may contribute to increased inflammation and pulmonary
hypertension (PH) in SCD patients (40). We show that treatment of THP-1, a
monocytic cell line, with ET-1 induced expression of a chemokine MIP-1β, a
potent chemoattractant for natural killer cells and T cells. ET-1 mediated MIP-1β
expression required the activation of PI-3 kinase, p38 MAP kinase, NADPH
oxidase, and N-terminal Jun kinase (JNK). In addition, our studies showed that
ET-1 induced MIP-1β expression required HIF-1α. The data was validated by the
use of siRNA for HIF-1α. Mutations in the five HREs present in the MIP-1β
proximal promoter attenuated ET-1 mediated MIP-1β promoter activity,
suggesting that all five HREs in the proximal promoter of MIP-1β were essential
for ET-1 mediated MIP-1β gene expression. We also investigated the possible
role of miRNAs in MIP-1β expression. Initial studies show decreased expression
levels for miRNAs- 20, 194 and 195. However, miR-223 showed increased
expression levels, suggesting a possible role for miRNAs-194 and 195, but not
85

miR-223 in ET-1 mediated MIP-1β expression. Further studies are warranted to
determine the role of miR-194 and miR-195 in MIP-1β regulation by anti-miR
approach.
In the third chapter, we examined the relevance of the data obtained in
vitro with that obtained in vivo in sickle cell disease. We determined the
leukotriene levels in transgenic sickle (TgSS) mice and patients with sickle cell
disease. Our studies showed that LTB
4
levels were 1.4 fold higher in the plasma
of TgSS mice compared to wild type (C57 B6) mice. Furthermore, analysis of
urine samples from sixty SCD patients showed ~ 2-fold increase in LTB
4
and
cysteinyl leukotrienes (Cyst LTs) levels compared to 15 healthy normal
volunteers. These results show that SCD patients have higher levels of
leukotrienes and inflammation, which could be attenuated by using
pharmacological inhibitors of HIF-1α, thus providing new avenues for therapeutic
treatment of inflammation, asthma, and vaso-occlusion in SCD patients.

86

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Creator Gonsalves, Caryn Suzanne (author)

Core Title A novel role for hypoxia-inducible factor-1alpha (HIF-1alpha) in the regulation of inflammatory chemokines and leukotriene expression in sickle cell disease

School Keck School of Medicine

Degree Doctor of Philosophy

Degree Program Biochemistry and Molecular Biology

Degree Conferral Date 2009-12

Publication Date 08/20/2009

Defense Date 06/25/2009

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag leukotrienes,OAI-PMH Harvest,sickle cell disease

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Kalra, Vijay K. (committee chair), Farley, Robert A. (committee member), Tahara, Stanley M. (committee member)

Creator Email beatrice_car2000@yahoo.co.in,caryn_g@hotmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-m2571

Unique identifier UC1173525

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Legacy Identifier etd-Gonsalves-2979.pdf

Dmrecord 252837

Document Type Dissertation

Rights Gonsalves, Caryn Suzanne

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Repository Name Libraries, University of Southern California

Repository Location Los Angeles, California

Repository Email cisadmin@lib.usc.edu