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Transcriptional regulation of IFN-γ and PlGF in response to Epo and VEGF in erythroid cells

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Transcriptional regulation of IFN-γ and PlGF in response to Epo and VEGF in erythroid cells

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Content i

TRANSCRIPTIONAL REGULATION OF IFN-γ AND PlGF IN RESPONSE TO
EPO AND VEGF IN ERYTHROID CELLS

BY

Ruchika Jaisinghani

A Thesis Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
MASTER OF SCIENCE
(MOLECULAR MICROBIOLOGY AND IMMUNOLOGY)

MAY 2013

Copyright 2013 Ruchika Jaisinghani

ii

Acknowledgements

I owe my enduring debt of gratitude to Dr. Vijay Kalra, for his individual attention,
constant encouragement and support, immense help, planning and valuable
guidance throughout the course of my work. His valued suggestions, strict
supervision of progress, continued feedback and update of relevant information
by way of additional input have generated great confidence and interest in me in
this specific field of study.

I express my heartfelt gratitude to Dr. Stanley Tahara, for the immense support
that he has extended during the tenure of my work. His instructive and rigorous
requirements towards scientific research always drove me to be a serious
researcher along this journey. I wish to express sincere thanks to Dr. Keigo
Machida for serving on my committee. I want to thank Dr. Caryn Gonsalves, for
guiding me throughout my work in the laboratory and giving me access to all
required materials during the course of my study. I sincerely appreciate her
efforts for rendering me wholehearted help all the way to complete my work. I am
especially grateful to her for teaching me all the techniques. I am deeply indebted
to Rachit Kumar for his tremendous motivation to move forward and confront all
challenges confidently. Also, I would like to extend my thanks to my lab
members, Chen Li and Pranali Taskar, for supporting me in my research and
instilling team spirit within me.

I want to make a special mention of the unlimited support given to me by my
parents, Mr. Manoj and Nancy Jaisinghani and my siblings, Anjali and Devesh.
They inspired me to work in an organized and systematic manner and have been
pillars of strength and patience. For all that I have achieved, I owe it to them for
being there always and hence making this project a great success.
iii

List of Abbreviations
5-LO 5- Lipoxygenase
Act D actinomycin D
AP-1 activator protein-1
ATF3 activating transcription factor 3
cAMP cyclic adenosine monophosphate
ChIP chromatin immunoprecipitation
CREB cAMP response element-binding protein
C-TAD C-terminal transactivation domain
Dn dominant negative
Epo erythropoietin
ET-1 endothelin-1
ET-1 endothelin-1
FLAP 5-lipooxygenase activating protein
GAPDH glceraldehyde 3-phosphate dehydrogenase
HbAS sickle cell trait
HbSS sickle hemoglobin
HDAC histone deacetylase
HIF hypoxia inducible factor
HIF-1 α hypoxia inducible factor-1 α
HLH helix-loop-helix
HPMVEC human pulmonary microvascular endothelial cells
HPMVEC Human pulmonary microvascular endothelial cells
HRE hypoxia-response elements
ICAM-1 intercellular adhesion molecule-1
IFN-γ interferon-gamma
iv

IL-3 interleukin-3
IL-8 interleukin-8
IMDM Iscove’s modified Dulbecco’s medium
MAPK mitogen-activated protein kinase
MCP-1 monocyte chemotactic protein-1
MIP-1β macrophage inflammatory protein-1β
MNC mononuclear cells
ODD oxygen degradation domain
PAI-1 plasminogen activator inhibitor-1
PAS domain PER-ARNT-SIM domain
PHD2 prolyl hydroxylase 2
PHT pulmonary hypertension
PI3K phosphoinositide 3-kinase
PlGF placenta growth factor
qRT-PCR quantitative real-time reverse transcription-PCR
RBCs red blood cells
RPA RNase protection assay
SCD sickle cell disease
shRNA small hairpin RNA
siRNA small interfering RNA
SS sickle cell disease-homozygous sickle globin
SSRBCs sickle red blood cells
TGF transforming growth factor
TNF-α tumor necrosis factor-α
tPA tissue plasminogen activator
TSA tricostatin A
VCAM-1 vascular cell adhesion molecule-1
VEGF vascular endothelial growth factor
v

VHL von Hippel-Landau protein
WBCs white blood cells
WT wild type
vi

Table of Contents

ACKNOWLEDGEMENTS II
LIST OF ABBREVIATIONS III
TABLE OF FIGURES VIII
LIST OF TABLES X
ABSTRACT XI
CHAPTER 1: INTRODUCTION 1
A. PATHOPHYSIOLOGY OF SICKLE CELL DISEASE 1
B. TRANSCRIPTIONAL REGULATION OF PLACENTAL GROWTH FACTOR (PLGF) 4
C. THE HYPOXIA-INDUCIBLE FACTOR FAMILY OF TRANSCRIPTION FACTORS 8
D. VASCULAR ENDOTHELIAL GROWTH FACTOR 11
E. ERYTHROPOIETIN 13
F. INTERFERON-GAMMA (IFN-Γ) 14
G. ACTIVATING TRANSCRIPTION FACTOR (ATF3) 15
CHAPTER 2: MATERIALS AND METHODS 19
A. ERYTHOID CELL CULTURE 19
B. REAGENTS AND ANTIBODIES 19
C. PROMOTERS, OVER-EXPRESSION AND DELETION CONSTRUCTS 20
D. MRNA EXTRACTION AND ANALYSIS BY QRT-PCR 20
E. CHROMATIN IMMUNOPRECIPITATION (CHIP) ASSAY 21
F. TRANSIENT TRANSFECTIONS 23
G. PROTEIN EXTRACTION 24
H. WESTERN BLOT ANALYSIS 24
I. STATISTICAL ANALYSIS 24
CHAPTER 3: VEGF AND EPO-MEDIATED INTERFERON GAMMA (IFN-γ)
EXPRESSION IS TRANSCRIPTIONALLY REGULATED BY ATF3
TRANSCRIPTION FACTOR 25
A. HYPOTHESIS 1 25
B. SPECIFIC AIM 1 25
C. RESULTS 26
vii

CHAPTER 4: ERYTHROPOIETIN AND VEGF-MEDIATED EXPRESSION OF
PLACENTAL GROWTH FACTOR AND ITS REGULATION BY HIF-1α IN
ERYTHROID CELLS 41
A. HYPOTHESIS 2 41
B. SPECIFIC AIM 2 41
C. RESULTS 42
DISCUSSION 49
BIBLIOGRAPHY 55

viii

Table of Figures

Figure 1: Diagrammatic comparison of Normal RBC with Sickled RBC ................ 2

Figure 2: Pathophysiology of vaso-occlusions in sickle cell disease ..................... 3

Figure 3: Molecular mechanisms of PlGF-induced proangiogenic signaling ......... 6

Figure 4: Effect of PlGF on endothelial cells and monocytes ................................ 8

Figure 5: Regulation of HIF-1α protein by prolyl hydroxylation and
proteasomal degradation ...................................................................... 10

Figure 6: Ligands and Receptors of VEGF Family .............................................. 12

Figure 7: Spliced isoforms of ATF3 ..................................................................... 17

Figure 8: IFN-γ mRNA expression in K562 cells in response to Epo and VEGF . 26

Figure 9: ATF3 mRNA expression in response to Epo and VEGF ...................... 29

Figure 10: ATF3 regulates IFN-γ gene expression .............................................. 31

Figure 11: Schematics of IFN-γ promoters. ......................................................... 32

Figure 12: Effect of Epo and VEGF treatment on IFN-γ luciferase promoter
activity ................................................................................................. 32

Figure 13: ATF3 regulates IFN-γ promoter luciferase activity ............................. 35

Figure 14: ATF3 binding to IFN-γ promoter, mediated by EPO and VEGF in
chromatin ............................................................................................ 36

Figure 15: Chromatin remodeling with HDAC inhibitor, Trichostatin A (TSA) ...... 38

Figure 16: Epo-treatment of K562 cells modestly reduces stability of IFN-γ ....... 39

Figure 17: Epo/VEGF-induced expression of PlGF in K562 erythroid cells ......... 42

Figure 18: Schematic of PlGF promoter (-2649 to +1 bp) .................................... 43

Figure 19: ATF3 and HIF-1α regulates PlGF expression .................................... 44
ix

Figure 20: Effect of Epo and VEGF on PlGF transcriptional activity .................... 45

Figure 21: Chromatin Immunoprecipitation analysis showed HIF-1α binding to the
PlGF promoter in the native chromatin ............................................... 47

Figure 22: Chromatin remodeling with HDAC inhibitor, Trichostatin A (TSA) ...... 48

x

List of Tables

Table 1: Primers used in the study of Epo and VEGF-mediated PlGF and IFN-γ
expression.
............................................................................................................................
23

xi

Abstract

Sickle cell disease (SCD) is a genetic disorder, characterized by a
mutation within the β-globin chain of the hemoglobin molecule. The clinical
manifestations of SCD include hemolytic anemia, vaso-occlusive crises and
pulmonary hypertension (PHT). To date, there is no permanent treatment for the
disease. SCD patients develop inflammation, PHT and reactive airway disease.
Our laboratory findings have shown that levels of placenta growth factor (PlGF)
are elevated in SCD, and mediate its effect on endothelial cells and monocytes,
in the expression of cytochemokines, endothelin-1 and PAI-1 by activating HIF-
1α, independent of hypoxia. Moreover, it has been shown that plasma levels of
interferon-γ (IFN-γ) among other cytokines are increased. However, relatively
less is known how expression of PlGF and IFN-γ are regulated, as understanding
of cell signaling pathways may provide insight to reduce the expression of these
molecules, which contribute to inflammation and PHT in SCD. We examined the
effect of erythropoietin (Epo) and VEGF, molecules whose expression increases
during hypoxia, seen in SCD.

In the present work, we showed that Epo and VEGF upregulated the
expression of IFN-γ in K562 erythroid cell line. Furthermore, shRNA for HIF-1α
reduced IFN-γ expression while ATF3 shRNA augmented IFN-γ expression.
Overexpression of ATF3 attenuated IFN-γ expression. Promoter analysis of IFN-
γ utilizing IFN-γ promoter luciferase plasmid, and ChIP, showed occupancy of
ATF3 in the promoter region of IFN-γ. Our studies for the first time, to the best of
our knowledge, showed that both HIF-1α and ATF3 were involved in the IFN-γ
expression, where ATF3 acted as a negative regulator of IFN-γ expression in
erythroid cells.

Next, we examined the mechanism of Epo and VEGF-mediated PlGF
expression in erythroid cells. Our studies indicated that both Epo and VEGF up
xii

regulated the expression of PlGF in K562 cells. Moreover, we inferred that both
HIF-1α and ATF3 were involved in its expression, as shRNA for HIF-1α and
ATF3 reduced PlGF expression. In silico analysis of PlGF promoter revealed the
presence of seven HRE sites in its promoter (~2 Kb). Promoter analysis utilizing
wild type, full- length promoter luciferase construct and mutation of selected HRE
sites in the promoter further confirmed the role of HIF-1α in PlGF expression.
Chromatin immunoprecipitation analysis (ChIP) utilizing antibody to HIF-1α
established that HIF-1α was bound to the PlGF promoter in native chromatin to
up-regulate the expression of PlGF.

In conclusion, these results showed that Epo and VEGF mediated
expression of IFN-γ and PlGF in erythroid cells was transcriptionally regulated by
HIF-1α and ATF3. Interestingly, ATF3 acted as a repressor or negative regulator
of IFN-γ expression in erythroid cells, mediated by Epo and VEGF.

1

Chapter 1: Introduction

A. Pathophysiology of Sickle Cell Disease

Sickle Cell Disease is an inherited blood disorder seen most commonly
among people of African ancestry, and also found in people of Mediterranean
and Middle Eastern background. As a genetic disease, it is characterized by
mutation of the β-globin gene, wherein glutamate is replaced with valine at the
6th position of β-globin. This leads to distortion of the discoid RBC shape to
sickled RBC (SS RBCs). The latter are less deformable (flexible) compared with
normal RBC. When these SS RBCs (6 micron in diameter) pass through small
capillaries (3 micron in diameter), these cells are trapped and adhere to vascular
endothelium, contributing to vascular occlusion and ischemia (hypoxia).
Repeated sickling and oxygenation-deoxygenation damages RBC membranes
and shortens their life span, resulting in hemolysis and anemia. Chronic
hemolytic anemia, shortened life span of RBCs, and vascular occlusions are the
hallmarks of the disease (Perelman et al., 2003).

Activation of endothelial cells in SCD contributes to vascular occlusions
(Belcher et al., 2000), (Solovey et al., 1999), (Sultana et al., 1998), (Hebbel,
1997). Plasma levels of vascular endothelial growth factor (VEGF), one of the
most potent activators of the endothelium, is elevated in SCD, and increases
expression of intercellular adhesion molecule-1 (ICAM-1) and VCAM-1 on
endothelial cells (Lu et al., 1999), which contributes to increased adhesion of
sickle RBCs to endothelial cells.

2

Figure 1: Diagrammatic comparison of Normal RBC with Sickled RBC

In recent years considerable evidence has accumulated that sickle cell
disease is an inflammatory state. Sickle patients have elevated white blood
counts, activated granulocytes, monocytes, and endothelial cells, exhibit
enhanced expression of endothelial cell adhesion molecules, elevated cytokine
levels, and elevated acute phase reactants. It is unclear whether inflammation is
a primary response of the polymerization of sickle hemoglobin or a secondary
response to tissue injury or infection. Another question that arises is how does a
mutation in the beta globin gene that promotes anemia and enhanced
erythropoiesis lead to an inflammatory phenotype that is
proangiogenic? (Vercellotti, 2003)

Two reports in Blood journal posit that placenta growth factor, PlGF, an
angiogenic factor belonging to the vascular endothelial growth factor (VEGF)
family, can be the tie that binds enhanced erythropoiesis, inflammation, and
angiogenesis together in sickle cell disease (Perelman et al., 2003). Studies
3

from our lab show that plasma PlGF levels are high in SCD and correlate with
sickle cell disease severity. Remarkably, PlGF is induced in bone marrow CD34
progenitor cells in the presence of erythropoietin (Epo). Thus, enhanced
erythropoiesis increases PlGF. In addition, PlGF expression leads to increases in
mRNAs of proinflammatory cytokines such as interleukin-1 (IL-1) and IL-8 as well
as VEGF itself; this was observed in monocytes obtained from normal individuals
and required signaling through VEGFR-1 (Selvaraj et al., 2003).

Figure 2: Pathophysiology of vaso-occlusions in sickle cell disease
Due to anemia, increased erythropoiesis occurs in SCD leading to
production of erythropoietin. Epo activates erythroid progenitor cells to produce
PlGF. The latter activates both monocytes and endothelial cells. Activation of
monocytes by PlGF leads to expression of cytochemokines (e.g. IL-1β, MCP-1).
These cytokines activate endothelium to upregulate the expression of adhesion
molecules (e.g. ICAM-1 and VCAM-1), the latter are involved in the adhesion of
SS RBC to endothelium leading to vaso-occlusion. This leads to hypoxia, which
in turn results in more sickling of red blood cells. This vicious cycle leads to
enhanced vaso-occlusion and inflammation. (Perelman et al., 2003)

4

PlGF is a potent stimulant for VEGF secretion by monocytes (Bottomley
et al., 2000) and is co-expressed with VEGF in synovial fluid where high levels of
PlGF homodimers and PlGF/VEGF heterodimers are found, suggesting that it
plays a role in the inflammatory process (Bottomley et al., 2000). Since
erythropoiesis is expanded in individuals with SCD to compensate for increased
hemolysis of SS RBC, the plasma levels of erythropoietin and VEGF are higher
(Solovey et al., 1997). Moreover, studies showed that that plasma levels of PlGF
are higher in SCD patients compared to matched normal individuals. It was
hypothesized that PlGF is the intrinsic red cell factor that mediates the activation
of leukocytes in SCD, contributing to vaso-occlusive crises (Tordjman 2001,
Perelman 2003). Nonetheless, the molecular mechanism of erythropoietin (Epo)
and VEGF mediated expression of PlGF in erythroid cells is not understood.
Furthermore, it is not clear how these molecules affect the expression of
inflammatory molecules, specifically interferon-gamma (IFN-γ).

B. Transcriptional Regulation of Placental Growth Factor (PlGF)

PlGF was originally cloned in 1991 from a cDNA library derived from
human term placenta. PlGF is highly expressed by trophoblasts (Khaliq et al.,
1996) (Shore et al., 1997) and decidual natural killer cells (Hanna et al., 2006)
localized within the human-maternal fetal interface. PlGF is also detected in other
tissues such as heart, lung, muscle and adipose tissue. PlGF belongs to the
vascular endothelial growth factor (VEGF) family of proteins, and there is a
sequence similarity of about 50% between VEGF A and PlGF.

PlGF is a dimeric protein and interstrand disulfide bonds hold monomers
together. Each PlGF monomer has eight cysteine residues that are engaged to
form three intra-chain disulfide bonds, generating a 3D structure known as a
cysteine-knot motif (De Falco, 2012). Additionally, PlGF is able to form
5

heterodimers with VEGF, especially when these two proteins are expressed in
the same cell (Bobic et al., 2012).

The human PlGF gene maps to chromosome 14q24, whereas the mouse
gene is located on chromosome 12qD. Both genes comprise 7 exons spanning
13.7 kb in human and 10.4 kb in mouse (Maglione et al., 1993). Due to
alternative splicing, different isoforms are encoded by the human PlGF gene. The
four PlGF isoforms (PlGF 1-4; (Maglione et al., 1993), comprise 131, 152, 203
and 224 amino acids respectively, after removal of the signal peptide.

The primary difference between the four isoforms is that PlGF-1 and
PlGF-3 are non-heparin binding diffusible isoforms, while PlGF-2 and PlGF-4
each have an additional heparin binding domain (Yang et al., 2003). By
comparison, mouse PlGF gene encodes a single isoform: PLGF-2, which is able
to bind heparin, and is comprised of 140 amino acids in its mature form (DiPalma
et al., 1996).

PlGF binds vascular endothelial growth factor receptor-1 (VEGFR1, or Flt-
1, i.e., Fms-like tyrosine kinase-1), but not VEGFR2, which is the main receptor
for VEGF-A signaling (Sawano et al., 1996). VEGFR1 is a receptor tyrosine
kinase, but is also expressed as a soluble molecule which, lacks transmembrane
and intra-cellular kinase domains; it acts as a decoy receptor for VEGF-A and
PlGF (Loges et al., 2009). Human cells of placental origin, called trophoblasts,
express VEGFR1, which is activated upon PlGF binding. In human umbilical vein
endothelial cells. PlGF binding results in release of nitric oxide and vasodilation
in uterine arteries. VEGFR1 is also expressed on monocytes, neutrophils,
eosinophils and smooth muscle cells (Bobic et al., 2012). The expression of
VEGFR1 is directly regulated by hypoxia and hypoxia-inducing factors (Bobic et
al., 2012).
6

Figure 3: Molecular mechanisms of PlGF-induced proangiogenic signaling
(A) VEGFA (VEGF) binds with a higher affinity to VEGFR-1 (designated
Flt1 in the mouse) than to VEGFR-2 (designated Flk2 in the mouse). Hence,
under physiological conditions, when PlGF expression is minimal, Flt1 acts as a
VEGF trap to prevent excessive VEGFR-2 activation. In cancer, PlGF is
upregulated in cancer cells and stromal cells, which leads to displacement of
VEGF from VEGFR-1. As a result, VEGF binds to VEGFR-2, leading to
transmission of a proangiogenic signal. (B) PlGF transduces its own signal via
VEGFR-1, as shown by phosphorylation of distinct tyrosine residues in the
intracellular domain of VEGFR-1. This signal leads to proliferation and migration
of tumor cells, endothelial cells, monocytes, and endothelial progenitor cells
(EPCs). Moreover, stimulation of VEGFR-1 by PlGF amplifies the VEGF-induced
signaling of VEGFR-2 by transphosphorylation of tyrosine residues of VEGFR-2.
As a result, VEGF-induced angiogenesis, vascular permeability and proliferation
are enhanced (Loges et al., 2009).

7

PlGF in circulation activates monocytes to synthesize cytochemokines,
which contributes to inflammation (Perelman et al., 2003). Studies show that
PlGF activates hypoxia inducible factor-1 α (HIF-1 α), in a hypoxia independent
manner, and regulates the expression of endothelin-1 from primary human
pulmonary microvascular cells (HPMVEC) and endothelin-B receptor in
monocytes (Patel et al., 2008). Subsequently, it was seen that PlGF activates
monocytes, leading to increased levels of leukotrienes, which contribute to lung
injury, i.e. reactive airway disease seen in SCD (Patel et al., 2009). Furthermore,
PlGF induction leads to increased levels of plasminogen activator inhibitor-1
(PAI-1), a primary inhibitor of fibrinolysis, in HPMVEC through activation of
transcription factors, HIF-1 α and activator protein-1 (AP-1) complex. These
findings were confirmed in vivo using genetic sickle mouse models, in mice with
knockdown of PlGF (PlGF-/-) and in normal mice over-expressing PlGF (PlGF
+/+). These studies showed that sickle mice express high levels of PlGF and
PAI-1. PlGF-/- mice, as expected, have low plasma PlGF. Conversely; PlGF +/+
mice exhibit high levels of plasma PlGF and increased levels of PAI-1. Thus, this
study provided an important lead towards understanding the role of elevated
levels of PlGF and PAI-1 in the development of hyper coagulation and pulmonary
fibrosis in SCD (Nitin Patel et al., 2010)

8

Bone Marrow Progenitor Cells

Erythroid Cells Mononuclear Cells

PlGF  VEGF 

PlGF causes the activation of Endothelial cells and Monocytes

PAI-1  Cytochemokines  Leukotrienes  ET-1 
MCP-1, IL-6, IL-8, (5-LO, FLAP)
TNF- α, IL-1 β)

Hyper coagulation Inflammation in Lung Injury in Vasoconstriction
Pulmonary fibrosis SCD SCD PHT in SCD

Figure 4: Effect of PlGF on endothelial cells and monocytes

C. The Hypoxia-inducible Factor Family of Transcription Factors

Oxygen gradients play an important and beneficial role in mammalian
physiology; adequate oxygen levels are necessary for various critical processes.
Therefore, cellular and systemic oxygen concentrations are tightly regulated via
pathways that affect the expression and activity of a number of cellular proteins.

Binds
to
VEGFR-‐1

Activates
HIF1-‐α
Activates
HIF1-‐α
Activates
HIF1-‐α

9

Hypoxia arises when demand for oxygen exceeds the supply. Low oxygen
tension or hypoxia provides the required extracellular stimulus for proper
embryogenesis and wound healing, and maintains the pluripotency of stem cells.
Hypoxia that involves oxygen tensions below the normal physiological range can
restrict the function of organs, tissues, or cells. Pathological hypoxia can be
caused by a reduction in oxygen supply, such as caused by high altitude or
localized ischemia due to the disruption of blood flow to a given area
(Pouyssegur et al., 2006). Extensive studies have been conducted on the oxygen
dependent induction of genes, such as the genes encoding erythropoietin (Epo),
VEGF (Wenger, 2000), (Semenza, 2002) and PlGF (Vercellotti, 2003).

The main effectors of hypoxic stimulus are the transcriptional factors
known as hypoxia inducible factors or HIF (Semenza, 1999). The HIF complex is
active only as a heterodimeric protein, with two subunits, the α subunit (HIF-1 α,
HIF-2 α or HIF-3 α) and the β subunit (HIF-1β, also known as aryl hydrocarbon
receptor nuclear translocator or ARNT), which together forms the HIF-1, HIF-2
and HIF-3 transcriptional complex (Wang et al., 1995). The HIF-1 α subunit is
regulated by oxygen tension, whereas the HIF-1 βsubunit is constitutively active
(Koh et al., 2012).

Under normal oxygen tension, HIF-1 α subunit undergoes hydroxylation
by prolyl hydroxylase-2 (PHD-2), and then undergoes degradation by the
ubiquitinin-ligase-proteosomal pathway. Thus basal levels of HIF-1 α protein are
low under normoxic condition. However, under hypoxia, PHD-2 does not carry
out hydroxylation of HIF-1 α, and thus levels of HIF-1 α are relatively higher
under hypoxia.

10

Figure 5: Regulation of HIF-1α protein by prolyl hydroxylation and
proteasomal degradation
There are three hydroxylation sites in the HIF-1 α subunit: two prolyl
residues in the oxygen-dependent degradation domain (ODDD) and one
asparaginyl residue in the C-terminal transactivation domain (C-TAD). In the
presence of oxygen, prolyl hydroxylation is catalyzed by the Fe (II)-, oxygen- and
2-oxoglutarate-dependent PHDs. The hydroxylated prolyl residues allow capture
of HIF-1 α by the von Hippel–Lindau protein (pVHL), leading to ubiquitination and
subsequent proteasomal degradation. In the absence of hydroxylation due to
hypoxia or PHD inhibition, HIF-1 α translocates to the nucleus, heterodimerizes
with HIF-1 β and binds to hypoxia-response elements (HREs) in the regulatory
regions of target genes.

Once HIF-1α translocates into the nucleus it heterodimerises with HIF-1βb
and binds to the hypoxia response elements or HRE (RCGTG or RGCAC) sites
within the promoter region of genes such as erythropoietin (Semenza et al.,
1992), (Ferrara et al., 1992) vascular endothelial growth factor (VEGF) (Forsythe
et al., 1996) and interleukin-8 (IL-8) (Kim et al., 2006).

11

Since higher levels of PlGF in SCD patients contribute to inflammation,
pulmonary hypertension and reactive airway disease (Perelman et al., 2003),
(Patel et al., 2008), (Patel et al., 2009), (N. Patel et al., 2010) it will be important
to understand how expression of PlGF is regulated at transcriptional and post-
transcriptional level. This will allow us to design therapeutic modalities to lower
the level of PlGF in circulation of SCD patients. Furthermore, it has been
observed that levels of interferon-gamma (IFN-γ) are high in SCD patients, which
contribute to inflammation (Wallace et al., 2009). However, relatively less is
known how IFN-γ levels are regulated in SCD patients.

D. Vascular Endothelial Growth Factor

Vascular Endothelial growth factor (VEGF), also known as vascular
permeability factor, was described as an inducer of angiogenesis in a variety of
physiological and pathological processes (Clauss et al., 1996). There are two
phospotyrosine kinase receptors for VEGF: the fms-like tyrosine kinase Flt-1
(VEGFR-1) and the fetal liver kinase, Flk-1 (VEGFR-2) or its human homolog
KDR (kinase insert domain-containing receptor). (Clauss et al., 1996). In
particular, VEGF, referred to also as VEGF-A, is a major regulator of normal and
abnormal angiogenesis, including that associated with tumors and several
intraocular syndromes. (Ferrara et al., 2001). The term VEGF refers to a
collection of related protein isoforms derived from the same gene (Ferrara et al.,
1992), the most well studied being VEGF121, VEGF165 and VEGF189.

12

Figure 6: Ligands and Receptors of VEGF Family

Previous studies stated that monocytes, in contrast to the endothelium,
express only the VEGF receptor Flt-1 (VEGFR-1) (Leung et al., 1989). Both
VEGF and PLGF stimulate tissue factor production and chemotaxis in monocytes
at equivalent doses indicating that both VEGF and PLGF mediate signaling via
VEGFR-1. (Tischer et al., 1991) Endothelial cells express both the VEGFR-1 and
VEGFR-2, and produce more tissue factor upon stimulation with VEGF than after
stimulation with PlGF. Neutralizing antibodies to VEGFR-2 reduce the VEGF-
stimulated tissue factor induction, but do not affect PlGF-induced tissue factor
induction in endothelial cells (Keck et al., 1989). Thus these studies indicate
VEGF mediated signaling involves VEGFR2 in endothelial cells, while PlGF
mediated signaling involved VEGFR1 (Clauss et al., 1996).

13

E. Erythropoietin

Erythropoietin (Epo) is a primary cytokine required for the development of
red blood cells, specifically driving definitive erythropoiesis (Wu et al., 1995). It is
a 34.4 kDa glycoprotein (Sanchez-Elsner et al., 2004). Reduced levels of oxygen
positively regulate the expression of the Epo gene. Erythropoiesis normally is
operating at a low basal level to compensate for the loss of old red cells;
however, in a situation of increased erythrocyte demand, such as low oxygen
tension, anemia or bleeding, the production of Epo is induced up to 1000-fold
above the normal (basal) levels (Ebert et al., 1999).

Epo is produced primarily by fetal liver (Zanjani et al., 1977) and adult
kidney (Jacobson et al., 1957), and to some extent by hematopoietic progenitor
cells (Stopka et al., 1998). Erythropoietin primarily affects red blood cell
progenitors and precursors (which are found in the human bone marrow) by
promoting their survival, inducing the globin genes and by inhibiting apoptosis.
Epo functions by binding to and activating the Epo receptor (EpoR), expressed
on the surface of committed erythroid progenitor cells. This in turn induces
erythroid progenitor cell survival, proliferation, and differentiation into circulating
enucleated hemoglobin-containing red blood cells (RBCs), which are critical for
oxygen transport. (Elliott et al., 2012).

Human Epo (HuEpo) is encoded by a single gene on chromosome 7 (Law
et al., 1986) and and on chromosome 5 in the mouse, that is transcribed into a
1.6–2.0 kb mRNA (Jacobs et al., 1985), and translated into a 193 amino acid
precursor protein. During transit through the secretory apparatus, the 27 amino
acid signal peptide and C-terminal arginine are removed, carbohydrate chains
are added (three N-linked and one O-linked) and the 30-kDa glycoprotein is
released into the surrounding fluids. The normal level of circulating Epo in
humans is approximately 5 pM (20 mU/mL; 100 pg/mL).
14

Previous studies showed Epo increased PlGF expression in Ewing
Sarcoma cells by activating metal transcription factor-1 (MTF-1) (Perelman et al.,
2003). Studies of Nishimoto et al., (Nishimoto et al., 2009) implicated MTF-1 in
the transcriptional regulation of PlGF in tropoblast-derived cells. Since the
promoter of PlGF has cis-binding elements for HIF-1α, PPAR-α and ATF3 in
addition to MTF-1, we examined their role in transcription of PlGF, mediated by
Epo and VEGF in erythroid cells.

F. Interferon-gamma (IFN-γ)

Interferon-gamma (IFN-γ) is a dimerized soluble cytokine that is the only
member of the type II class of interferon. IFN- γ is produced by natural killer (NK),
NKT, CD4+ and/or CD8+ T cells stimulated by the T-helper cytokines IL-12 and
IL-18 (Schroder et al., 2004). This interferon was later called macrophage-
activating factor, a term now used to describe a larger family of proteins to which
IFN- γ belongs. In humans, the IFN- γ protein is encoded by the IFNG gene
(Naylor et al., 1983).

IFN-γ functions at the innate-adaptive immune interface to promote
antibacterial, antitumor and antiviral responses. It acts as central mediator of
protective immune responses against the pre-erythrocytic and blood cell stages
of malaria (McCall et al., 2010). It also modulates fetal hemoglobin synthesis in
sickle cell anemia and thalassemia (Miller et al., 1990). Erythroid cells, under
acute hypoxia or when treated with phenyl hydrazine, showed the presence of
IFN-γ mRNA. Moreover, after cell cultivation with erythropoietin, qualitative and
quantitative changes in gene expression of cytokines were observed (Kozlov et
al., 2001).

15

It was shown that human CD4+ T cells produce increased levels of IFN- γ
in the presence of IFN- α , which suggests that type-1 IFNs favor Th1
differentiation (Brinkmann et al., 1993). Furthermore, IFN- α was shown to
increase IFN- γ mRNA synthesis in human T cells (Sareneva et al., 1998).
Although much is known about the regulation of IFN- γ expression in T cells, less
is understood about this process in erythroid cells.

G. Activating Transcription Factor (ATF3)

Activating transcription factor (ATF3) is a member of the ATF/CREB family
of basic leucine zipper-type (bZIP) transcription factors. ATF/cyclic AMP
response element-binding (CREB) family members include ATF1, CREB, ATF2,
ATF3, ATF4, ATF5, ATF6, ATF7 and B-ATF. The common feature that these
proteins share is the bZIP element (Thompson et al., 2009).

ATF3 is composed of 181 amino acids with a calculated mass of 22 kD.
The basic region and leucine zipper domain from amino acids 85 to 147 are
required for dimer formation and specific DNA binding (Hsu et al., 1992) (Chen et
al., 1994). The basic region in this domain is responsible for specific DNA
binding, while the leucine zipper region is responsible for forming homodimers or
heterodimers with other bZIP containing proteins such as the AP-1 or Maf
families of proteins or other ATF/CREB proteins, including ATF2 (T. W. Hai et al.,
1989), c-JUN (T. Hai et al., 1991) and JUN B (Hsu et al., 1992). The homodimer
of ATF3 represses transcription from various promoters with ATF sites, whereas
heterodimers with c-Jun or JunB activate transcription (Hua et al., 2006). Thus,
ATF3 can act as a repressor or activator of transcription.

16

In addition to the heteromeric complexity, various spliced isoforms of
ATF3 may further generate functional diversity in different cellular contexts (Hua
et al., 2006). We used two different variants of overexpressing ATF3, Variant 1
and Variant 4. Variant 1 encodes the longest isoform (1) of ATF3, which
homodimerizes and represses transcription from promoters with ATF binding
elements. Variant 4 (also known as ΔZip2a) contains an alternate terminal exon,
thus differing in the 3 ’ coding regions and 3 ’UTR from Variant 1. The encoded
isoform 2 (also known as Δ Zip2) has a distinct and shorter C-terminus,
compared to isoform 1. Isoform 2 lacks the leucine zipper protein-dimerization
motif and is capable of DNA binding and stimulates transcription, presumably by
sequestering inhibitory co-factors away from the promoter, thus counteracting the
transcriptional repression by full-length ATF3 by acting dominant negative.
(Hashimoto et al., 2002).

17

(Activating Transcription Factor 3) ATF3

ATF3 ΔZip ATF3 ΔZip2 ATF3 ΔZip3
Isolated in serum- lacks leucine zipper domain identified in amino
stimulated HeLacells. and thus is capable of DNA acid derived cells.
binding. It is localized in the
nucleus and counteracts the ATF3 ΔZip3b
transcriptional regulation by It is implicated in
full-length ATF3. mediating cAMP si-
gnalling of progluc-
agon transcription
in pancreatic α -
cells

ATF3 ΔZip2a and –b ATF3 ΔZip2c
1. Isolated from cells treated with various identified in amino acid
stimuli such as A23187, TNF- α, endopl- derived cells.
asmic reticulum stress or oxidative stress.
2. These isoforms encode the C-terminally
truncated protein of 135 amino acids,
which shares the N-terminal 116 amino acids
with the full-length ATF3 but contains novel
19 amino acid at the C-terminus.

Figure 7: Spliced isoforms of ATF3

18

ATF3 is induced upon exposure of cells to a variety of physiological and
pathological stimuli. It is required to for regulation and for limiting key aspects of
immune function, particularly in preventing immune pathologies associated with
uncontrolled pro-inflammatory cytokine production (Hashimoto et al., 2002;
Thompson et al., 2009). ATF3 deficiency is detrimental to inflammatory disease
conditions, including septic shock and asthma. Conversely ATF3 deficiency has
been shown to be beneficial for the clearance of MCMV infection by modulating
the IFN γexpression in natural killer cells (Rosenberger et al., 2008). ATF3 acts
as an important negative gene regulator of pro-inflammatory cytokine gene
expression (IFN- γ) in macrophages (Gilchrist et al., 2006). On the other hand it
has been shown that ectopic expression of ATF3 in CD4(+) T cells enhanced the
production of IFN-γ, the hallmark cytokine of TH1 cell, whereas small interfering
RNA knockdown of ATF3 reduced the IFN-γ production. Hence, ATF3 can both
positively and negatively regulate IFN-γ gene expression (Filen et al., 2010).
Furthermore, it has been shown recently that ATF3 is a target of the proto-
oncogene c-myc in serum induced-cell proliferation (Tamura et al., 2005).

As the role of ATF3 varies within cells, it is therefore, important to identify
ATF3 transcriptional targets and interacting partners in order to accurately gauge
the contribution of ATF3 to transcriptional regulation within a particular cell line.
Hence, I am examining the role ATF3 plays as a potential positive or negative
regulator of PlGF and IFN-γ inresponse to EPO and VEGF, in erythroid cells.
Based on preliminary data, I hypothesize that Epo and VEGF-mediated IFN-γ
expression may be negatively regulated by ATF3 in K562 cells, which is a human
erythroid cell line.

19

Chapter 2: Materials and Methods

A. Erythoid Cell Culture

K562 cells were the first human immortalized myelogenous leukemia cell
line. These erythroleukemia type cells (obtained from American Type Cell
Culture, or ATCC) were maintained in Iscove’s modified Dulbecco’s medium
(IMDM, Cellgro, Manassas, VA), supplemented with 10% heat inactivated fetal
bovine serum (FBS), 1mM L-glutamine, and 1X penicillin-streptomycin, and
were cultured at 37°C in an atmosphere of 5% CO2 in a humidified tissue culture
incubator. All experiments were performed using cells in the exponential growth
phase. Cells were kept in serum free media overnight prior to stimulations.

B. Reagents and Antibodies

Erythropoietin (Epo) was a kind gift from Dr. Vinod Pullarkat (Keck School
of Medicine, University of Southern California, Los Angeles, CA). Vascular
Endothelial Growth Factor (VEGF) was purchased from Peprotech (New Jersey,
US). Actinomycin D (Act D) is a most significant member of actinomycines,
which are a class of polypeptide antibiotics that has the ability to inhibit
transcription and was obtained from Sigma-Aldrich (St. Louis, MO). Trichostatin
A (TSA), an organic compound that serves as an antifungal antibiotic and
selectively inhibits the class I and II mammalian histone deacetylase (HDAC)
families of enzymes, but not class III HDACs was generously provided by Dr.
Shuping Zhong (Keck School of Medicine, USC, Los Angeles, CA). Primary
antibodies for Hypoxia Inducible Factor (HIF1 α), Activating Transcription Factor
3 (ATF3) and HRP-conjugated secondary antibodies were purchased from Santa
Cruz Biotechnology, Inc (Santa Cruz, CA). The β -actin conjugated HRP
20

antibody was purchased from Sigma-Aldrich (St. Louis, MO). Unless otherwise
specified, all other reagents were purchased from Sigma-Aldrich (St. Louis, MO).

C. Promoters, Over-expression and Deletion Constructs

PlGF promoter construct was synthesized in Dr. Kalra’s laboratory. The
pIFNG 1147-GL4.10 full-length IFN- γ promoter construct and the deletion
constructs (pIFNG1147-del2/3/4/5-GL4.10, pIFNG1147-del1/3/4/5-GL4.10,
pIFHG1147-del1/2/4/5-GL4.10, pIFNG1147-del1/2/3/5-GL4.10, pIFNG1147-
del1/2/3/4-GL4.10, pIFNG1147-del1/2/3/4/5-GL4.10) were a kind gift from Dr.
Riitta Lahesmaa (Turku Centre of Biotechnology, University of Turku,
Finland)(Filen et al., 2010). pGL3 and pGL4 constructs was obtained from
Promega Corporation (Madison, WI). Control shRNA (pGzip) and shRNA for
ATF3, HIF1- α, NRF1, NRF2, MTF-1 were generously provided by Dr. Stanley
Tahara (University of Southern California, Los Angeles, CA), obtained originally
from OpenBiosystems. ATF3 Over-expressing variants 1 and 4 were purchased
from OriGene Technologies, Inc (Rockville, MD).

D. mRNA Extraction and Analysis by qRT-PCR

K562 cells were treated with Epo (3 units/ml) and VEGF (250 ng/ml) for
indicated time periods followed by total mRNA extraction using TriZol (Invitrogen,
Carlsbad, CA). Real-time quantitative PCR of PlGF, IFN- γ and GAPDH were
performed using the iScript SYBR Green One-Step RT-PCR Kit (Bio-Rad,
Hercules, CA), using specific primers as listed in Table 1. Real time PCR
analyses were performed at the Analytical-Metabolic-Instrumentation Core of the
USC Research Center for Liver Disease (NIH grant P30 DK048522).
21

PCR amplification were carried out using 100 ng/µl of RNA for 40 cycles
under the following conditions: cDNA synthesis at 50°C for 10 min, iScript
reverse transcriptase inactivation at 95°C for 5 min, PCR cycling and detection at
95°C for 10s, and followed by elongation at 60°C for 30s, utilizing the ABI 7900
HT sequencing detection system (Life Technologies, Carlsbad, CA). Values were
expressed as relative expression levels of mRNA normalized to housekeeping
GAPDH mRNA levels. Relative quantification (RQ) values for mRNA expression
were calculated as 2
- ΔΔCt
by the comparative Ct method (Pfaffl, 2001), where Δ
ΔCt = (Ct target gene of treated sample – Ct GAPDH of treated sample) - (Ct
target gene of control sample - Ct GAPDH of control sample).

E. Chromatin Immunoprecipitation (ChIP) Assay

K562 cells were kept overnight in serum-free media, followed by treatment
with Epo and VEGF for the indicated time periods. ChIP analysis was performed
utilizing either HIF-1 α antibody or ATF3 antibody. Briefly, cells were cross-linked
with formaldehyde followed by treatment with, 1 M glycine to stop the cross-
linking. Cells were lysed and chromatin was sheared by sonication through
sonicator (6 pulses at 15 sec each, 40% efficiency/potency). The lysate was
centrifuged at 12,000 rpm for 10 min at 4°C using a microcentrifuge. The
supernatants were pre-cleared for 2 hr at 4°C with Protein A-Sepharose beads
(Sigma-Aldrich, St.Louis, MO) to block the non-specific interactions. Pre-cleared
supernatants were immune-precipitated with 1 μg of either HIF-1 α antibody,
ATF3 antibody, or control normal rabbit IgG antibody overnight at 4°C. 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°C overnight, and DNA was extracted by
phenol/chloroform/isoamyl alcohol followed by ethanol precipitation. Immuno-
22

precipitated 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°C for 30s, 60°C for 60s and 72°C for 120s, using primers listed in
Table 1. The PCR products were subjected to agarose gel electrophoresis and
examined for the binding of HIF1 α and ATF3 to thePlGF promoter and IFN γ
promoter, respectively. The values were normalized to input DNA.

Gene Method Forward Primer Reverse Primer
PlGF mRNA qRT-
PCR
5’-tgttcagcccatcctgtgtc-3’ 5’-acagtgcagattctcatcgcc-3’
ATF3 mRNA qRT-
PCR
5'-tagcattacgtcagcctggg-3' 5'-agcgttgcatcacccctttt-3'
GAPDH
mRNA
qRT-
PCR
5'-aacctgccaagtacgatgacatc-
3'
5'- gtagcccaggatgcccttga-3'
IFN-γ mRNA qRT-
PCR
5'ctcgaaacagcatctgactcctt-3' 5’tgtccaacgcaaagcaataca-3’
PlGF
promoter
(HRE at -937/
933)
ChIP 5'-ggacacagaaggcag-3' 5'-tctgtccgctgtgtat-3'
PlGF
promoter
(HRE at -
1461/1457)
ChIP 5'-cacatcccagcataagtgc- 3' 5'-tgcatgtgtgtgagggtgagt-3'
PlGF
promoter
(HRE at -
1744/1741)
ChIP 5'-ggccaacatgacaaaaacctg-
3'
5'- gcacttatgcatgggatgtg-3'
IFN-γ
promoter
(ATF3 at -
62/52)
ChIP 5’-aggaggtgcagcacattgtt-3’ 5’-atgggtcctggcagtaacag-3’
IFN-γ
promoter
(ATF3 and
ChIP 5’-cgaagtggggaggtacaaaa-3’ 5’-gtgacagataggcagggatga-
3’
23

JUN B at -
200/-52)
IFN-γ
promoter
(ATF3 at -
1925/ -1918)
ChiP 5’-acgaggtcaggagatcgaga-3’ 5’-gacggagtcttgctctgtca-3’

Table 1: Primers used in the study of Epo and VEGF-mediated PlGF and
IFN-γ expression.

F. Transient Transfections

K562 cells (1 x 10
6
cells) were plated in 60 mm dishes containing 2 ml of
complete media containing FBS and antibiotics. K562 cells were transfected with
indicated hRNA constructs (1 µg). For reporter luciferase assay, 1 µg of either
PlGF-reporter or IFN-γ luciferase reporter constructs was co-transfected with 0.5
µg of β-galactosidase construct using HiPerFect Transfection Reagent (Qiagen,
Valencia, CA). Briefly, 6 µl HiPerFect Transfection Reagent was added to 100 µl
of culture medium without serum, and allowed to stand for 5-10 mins. 1 µg of
shRNA, promoter luciferase construct or over-expression plasmid was added to
the transfection mix and incubated for 20 minutes at room temperature to allow
the formation of transfection complexes. The complexes were added to the cells.
Transfected cells were kept in serum free media for a minimum of 3 hours or a
maximum of 24 hrs, and then stimulated with either Epo or VEGF for indicated
time periods. For luciferase assays, cells were harvested and analyzed for
luciferase activity (Promega, Madison, WI) using a luminometer (Berthold
Technologies; Lumat LB 9501, Germany). The light emitted during the initial 20 s
of the reaction, being linear, was measured. 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. For mRNA analysis, cells were lysed in TRIzol and mRNA isolated as
described above.
24

G. Protein Extraction

Approximately 5 x 10
6
K562 cells were washed in 1 ml of PBS and re-
suspended in 100 μl of RIPA lysis buffer (5 M NaCl, 500 mM EDTA, 500 mM
EGTA, 10% SDS, 10 mg/ml PMSF, 100% Nonidet P-40, 1 M Tris (pH 8.0) and
1X protease inhibitor mixture) and left in -80° for 30 minutes to swell. The lysate
was thawed and centrifuged at 12,000 rpm for 10 minutes. Supernatants (about
100 μ l) were collected as protein extracts. Protein concentrations were
determined using the Bradford method. (Bradford, 1976)

H. Western Blot Analysis

Protein extracts were used to determine ATF3 protein levels. 100 ng of
protein were run on an SDS-PAGE gel and were transferred to nitrocellulose
membrane. Membranes were blocked with 5% non-fat milk. They were probed
using an antibody specific for ATF3 (1:500). These membranes were then
stripped with stripping buffer (Bioland, Paramount, CA) and re-probed for β-actin
(1:2500) levels to determine equal loading of protein. Develop the membrane
using West Pico-ECL substrate, an enhanced chemiluminescent reagent, for
detection of horseradish peroxidase (HRP) activity, obtained from Thermo
Scientific Pierce Biotechnology, Inc (Rockford, IL).

I. Statistical Analysis

Results are expressed as mean ± S.E. The significance of differences in
mean values between untreated and VEGF/Epo-treated samples was
determined by Student's t test. Results were considered statistically significant at
p<0.05.
25

Chapter 3: VEGF and Epo-mediated Interferon gamma
(IFN-γ) Expression is Transcriptionally Regulated by
ATF3 Transcription Factor

A. Hypothesis 1

It has been shown that higher levels of IFN-γ are present in plasma of
SCD patients compared to normal individuals (Taylor et al., 1990). However, it is
yet to be understood how the expression of this molecule is regulated. Our
preliminary studies showed treatment of a human erythroid cell line K562, with
Epo augmented the expression of IFN-γ. The promoter of IFN-γ has cis-
consensus binding elements for ATF3 in addition to JUN and AP-1. Furthermore,
our studies showed shRNA for ATF3 induced IFN-γ expression, indicating that
ATF3 may act as a negative regulator of IFN-γ. We hypothesize that ATF3 may
regulate the transcription of IFN-γ gene in erythroid cells. To address this, I have
developed the following specific aim.

B. Specific Aim 1

To determine whether ATF3 acts as a repressor of IFN-γ expression in
erythroid cells, in response to Epo and VEGF, I will utilize over-expressing ATF3
plasmids. Moreover, I will identify the role of Jun, AP-1 and ATF-3 in the
transcription of IFN-γ using IFN-γ promoter luciferase construct, and validate the
results utilizing chromatin immunoprecipitation (ChiP) analysis.

26

C. Results

1. Erythropoietin/VEGF mediated IFN-γ mRNA expression in K562 erythroid
cell line.

Figure 8: IFN-γ mRNA expression in K562 cells in response to Epo and
VEGF
(A) Epo-mediated IFN-γ expression was maximal at 1hr, while (B) VEGF
mediated IFN-γ expression was maximal at 2hr. At later time period (4-24 hrs),
both VEGF and IFN-γ mRNA expression showed a continuous decline.

27

Previous studies indicate that plasma levels of IFN-γ are high in SCD
patients compared to matched controls, thus we determined the expression of
IFN-γ in erythroid cells in response to erythropoietin (Epo) and Vascular
Endothelial Growth Factor (VEGF) (Taylor et al., 1990). We used K562 erythroid
cells for ease of culture, though the data obtained will be validated in erythroid
cells obtained from normal and SCD patients. Treatment of K562 cells with Epo
(3 units/ml) showed time dependent increase in IFN-γ mRNA expression, with
maximal increase at 1hr (Figure 8A). Thus, all subsequent studies with Epo were
carried out at 1 hr post-induction. However, treatment with VEGF (250 ng/ml),
another cytokine, showed maximum expression of IFN-γ at 2 hr. (Figure 8B)
Thus studies with VEGF were carried out at 2 hrs post-induction.

2. Role of ATF3 transcription factor in the regulation of IFN- γ.

We examined the expression of ATF3 in erythroid cells in response to Epo
and VEGF. As shown in (Figure 9A), Epo treatment resulted in a time-dependent
increase in ATF3 levels, with maximal increase at 1 hr, the same time period at
which we saw maximal increase in IFN-γ expression. Similarly, VEGF mediated
ATF3 maximal expression coincided with IFN-γ maximal expression at 2 hr post -
induction (Figure 9B). We observed Epo-mediated ATF3 increased several-fold
(~25-fold) after 24 hr, which was associated with a several-fold decrease in IFN-γ
expression (Figure 9C). Moreover, ATF3 shRNA reduced ATF3 protein
expression while over-expression ATF3 plasmid increased ATF3 protein (Figure
9D).

28

29

Figure 9: ATF3 mRNA expression in response to Epo and VEGF
(A) and (B) The maximal time of Epo and VEGF mediated ATF3
expression coincided with IFN-γ mRNA gene expression. (C) Epo-mediated
ATF3 mRNA increased several fold (~25-fold) while IFN-γ expression was
attenuated several fold at 24hr time period, indicating a reciprocal relationship
between ATF3 and IFN-γ expression at 24hr time period. (D) K562 cells were
treated with Epo for 6hrs and 100µg of protein extracts were subjected to
western blotting. Bands were detected using an antibody to ATF3. The
membrane was stripped and reprobed with antibody to β-actin, to normalize
loading. Protein bands were detected by chemiluminescence. Western blot
analysis showed ATF3 shRNA reduced while overexpression ATF3 plasmid
increased ATF3 protein.

In silico analysis of IFN-γ promoter (~2 kb) revealed the presence of two
ATF3 and several other hypoxia-response elements (HRE) proximal to the
transcription start site. Recent studies have shown opposite roles of ATF3 in
IFN-γ expression. As evidence, differentiation of early Th1 cells with IL-12 or IL-
18 led to increased IFN-γ expression, which was positively regulated by ATF3
(Filen et al., 2010). Conversely, in another study, virally induced IFN-γ
expression was negatively regulated by ATF3 (Rosenberger et al., 2008). For
these reasons, I examined the effect of ATF3 shRNA on IFN-γ expression in
erythroid cells.

Transfection of K562 cells with ATF3 shRNA, followed by Epo treatment
for 1 hr showed ~20-fold increased expression of IFN-γ, while control shRNA had
no effect (Figure 10A). In parallel, the effect of shRNA ATF3 on IFN-γ expression
in erythroid cells when treated with VEGF was also examined. The latter
conditions showed ~10-fold increased expression of IFN-γ, while control shRNA
had no effect (Figure 10B). These data are consistent with ATF3 acting as a
repressor to negatively regulate IFN-γ expression in the basal cell state. To
further validate these results, K562 cells were transfected with ATF3 over-
expression plasmid. As shown in Figure 10C, the levels of IFN-γ were ~40% and
30

~80% reduced in response to overexpression of ATF3 variant 1 and variant 4,
respectively. Both variants of ATF3 showed similar activity with varying extents of
down-regulation (repression). This could be possible binding of Variant 4 to full-
length ATF3, hence counteracting its transcriptional activity, thus working as a
dominant negative. (Hua et al., 2006). Taken together, these data showed that
ATF3 acted as a negative regulator of IFN-γ gene expression in K562 erythroid
cells.

31

Figure 10: ATF3 regulates IFN-γ gene expression
(A) and (B) K562 cells were transfected with shRNA for ATF3 and HIF-1α
followed by the treatment with Epo and VEGF, as indicated in the figures for 1 hr
and 2 hrs respectively. Epo- and VEGF-mediated IFN-γ mRNA expression was
attenuated by shRNA for ATF3 and HIF-1α. (C) Over-expression of ATF3 (o/e
ATF3), utilizing variant 1 and variant 4, reduced IFN-γ expression, indicating
ATF3 acted as a negative regulator for IFN-γ expression.

3. Promoter analysis of IFN-γ promoter.

32

Figure 11: Schematics of IFN-γ promoters.
(A) In silico analysis of IFN-γ promoter (+1 to -2126) shows the cis-binding
consensus elements for transcription factors, ATF3, JUNB, CREB, HIF-1α
(hypoxia response element) and AP-1. (B) Fragment of wild type IFN-γ promoter
(-1147 to +38) containing one ATF3, two AP-1, one JUN B and one CREB
binding site. AP-1, JUN B and ATF3 are the deletion constructs of IFN-γ
promoter in which only AP-1, JUN B and ATF3 sites are conserved respectively.
In IFN-γ Δ, all the sites are deleted. Note that HRE are conserved in all the
deletion contructs.

Figure 12: Effect of Epo and VEGF treatment on IFN-γ luciferase promoter
activity
(A) and (B) K562 cells were co-transfected with IFN-γ reporter plasmids
along with deletion constructs and β-galctosidase plasmid, the latter for
33

normalizing transfection efficiency. The luciferase activity was corrected by
subtracting background activity observed for promoter less pGL4 vector. Cells
were transfected with wild type (wt, -1147 to +38) and deletion constructs in
which all sites except the one shown was intact. For example, deletion construct
with ATF3 intact had deletion in AP-1 and Jun B. It should be pointed out that
HRE sites were intact in all of these constructs. This was followed with the
treatment of Epo/VEGF for 6 hrs. Both luciferase and β-galctosidase activity was
measured as mentioned in ‘Materials and Methods’. The luciferase activity was
normalized to that of the untreated IFN-γ-luc construct. Data are expressed as
Mean± SEM of n=3.

As shown in Figure 11A, the IFN-γ promoter (~2kB) has several HRE,
JUNB, AP-1 and ATF3 binding sites. We obtained a fragment of full-length IFN-γ
promoter, (-1147 to +38 bp) luciferase construct which contains one ATF3, two
AP-1, one Jun B and one CREB binding site (Filen et al., 2010) (Figure 11B).
Transfection of K562 cells with wild type IFN-γ promoter followed by Epo/VEGF
treatment for 6 hrs showed 1.5-fold increase in luciferase activity (Figure 12A and
12B). However, the deletion construct in which JUNB and AP-1 binding sites
were deleted, but not the ATF3 sites, showed ~2-fold increase in luciferase
activity. Other promoter constructs, in which the AP-1 and JunB binding sites
were intact, also showed luciferase activity to the same extent as observed with
IFN-γ wt promoter plasmid. However deletion of all sites for ATF3, AP-1 and
JUNB reduced promoter activity to the basal level. Taken together, these results
suggested that ATF3, AP-1 and JUN B together, were involved in transcriptional
regulation of IFN-γ, in response to both Epo and VEGF (Figure 12A and 12B).
Thus ATF3 may act as a repressor of IFN-γ in combination with another
transcriptional factor. Further studies are ongoing to substantiate these results.

My results strongly suggest that ATF3 acts as a negative regulator of IFN-
γ gene expression. To further corroborate my results with an alternative
approach, I examined the effect of ATF3 shRNA on IFN-γ promoter in erythroid
cells. Transfection of K562 cells with ATF3 shRNA, followed by Epo/VEGF
treatment showed ~20-fold increased luciferase activity driven from IFN-γ
34

promoter, while control shRNA had no effect on the luciferase activity (Figure
13A and 13B). Additionally, IFN-γ promoter luciferase activity was measured with
cells overexpressing ATF3. As shown in Figure 13C, the luciferase activity
transcribed off IFN-γ promoter was ~40% reduced in response to overexpression
of ATF3 variant 4. Taken together, these results are consistent with ATF3 acting
as a negative regulator of IFN-γ gene transcription in erythroid cells.

35

Figure 13: ATF3 regulates IFN-γ promoter luciferase activity
K562 cells were co-transfected with IFN-γ reporter plasmid and β-
galctosidase plasmid, the latter for normalizing transfection efficiency. The
luciferase activity was corrected for nonspecific luciferase expression by
subtraction of activity observed with promoter less pGL4 vector. (A) and (B) Cells
were transfected with wild type (wt, -1147 to +38) and shATF3 followed by the
treatment with Epo/VEGF for 6hrs. ATF3 shRNA augmented Epo and VEGF
mediated IFN-γ promoter luciferase activity, whereas the scrambled shRNA
shows no change in its effect. (C) Cells were co-transfected with wt promoter-
luciferase reporter and ATF3 expression plasmids. Overexpression of ATF3
(variant 4) reduced luciferase activity compared to wt control. Both luciferase and
β-galctosidase activity were measured as described in ‘Materials and Methods’.
The luciferase activities were normalized to that of the untreated IFN-γ-luc
construct. Data are expressed as Mean± SEM of n=3.

4. Chromatin Immunoprecipitation (ChIP) with ATF3 antibody.

36

Figure 14: ATF3 binding to IFN-γ promoter, mediated by EPO and VEGF in
chromatin
(A), (C) and (E) Chromatin was isolated from Epo-treated cell for 1hr time
period and cells were immunoprepitated using antibody to ATF3 (middle panel)
or control rabbit IgG (lower panel). PCR was performed utilizing primers
corresponding to proximal ATF3 site, ATF3 and JUN B site and a distal ATF3
site. (D) (E) and (F) Chromatin was isolated from VEGF-treated cells for 2hrs
time period and cells were immunoprecipitated using antibody to ATF3 (middle
panel) or control rabbit IgG (lower panel). PCR was performed utilizing primers
corresponding to ATF3 site, ATF3 and JUN B site and a distal ATF3 site. The
data shows equal loading of input DNA (upper panel) and absence of expected
PCR product in IgG immunoprecipitated samples.
37

Chromatin Immunoprecipitation with ATF3 antibody was carried out to
check the binding efficiency of ATF3 to IFN-γ promoter. The data showed Epo
reduced the binding of ATF3 to the TSS-proximal site in the IFN-γ promoter,
however there was no change in the binding of ATF3 to combined site of ATF3
and JUN B and also to the distal site. (Figure 14A, 14C and 14E). Cells treated
with VEGF showed that ATF3 binds to the IFN-γ promoter at the distal ATF3 site
and showed further reduction in binding to the combined ATF3 and JUN B site
whereas the ATF3 site one was not affected. (Figure 14B, 14D and 14F) Taken
together, these data showed that Epo reduced ATF3 binding to the TSS -
proximal site in the IFN-γ promoter and thus likely augmented IFN-γ expression.
Conversely, VEGF reduced binding of ATF3 to the distal site of IFN-γ promoter
and thereby increasing IFN-γ expression. Moreover TSS-proximal ATF3 site is
likely not involved in VEGF-mediated IFN-γ expression. We utilized histone
deacetylase inhibitor TSA to gain insight whether chromatin remodeling was
involved in EPO-and VEGF-mediated IFN-γ expression. In chromatin derived
from TSA-treated erythroid cells there was reduced binding of ATF3 to the TSS-
proximal and combined ATF3 and JUN B sites but the data were inconclusive
and further studies are warranted for fully understanding the role of chromatin
remodeling in transcription of the IFN-γ gene.

5. Chromatin remodeling with HDAC inhibitor TSA.

Tricostatin A (TSA) is an organic compound that serves as an antifungal
antibiotic. It is used to alter gene expression by interfering with the removal of
acetyl groups from histones (histone deacetylases, HDAC) and therefore
alternating the ability of DNA transcription factors to access the DNA molecules
inside chromatin, hence promotes histone acetylation by opening up the
chromatin structure. Our studies showed TSA augmented IFN-γ expression
indicating chromatin remodeling allows upregulation of IFN-γ expression under
38

basal and VEGF treated conditions (Figure 15B), but no additive or synergistic
effect was observed with Epo. (Figure 15A)

Figure 15: Chromatin remodeling with HDAC inhibitor, Trichostatin A (TSA)
(A) and (B) TSA, an inhibitor of histone deacetylase, augmented IFN-γ
expression indicating chromatin remodeling allows upregulation of IFN-γ
expression under basal and VEGF treated condition, but no additive or
synergistic effect was observed with Epo.

39

6. Post-transcriptional regulation of Epo-mediated IFN-γ mRNA expression.

Figure 16: Epo-treatment of K562 cells modestly reduces stability of IFN-γ
K562 cells were treated with and without Epo for 1 hr, followed by
incubation with actinomycin D, a transcriptional inhibitor, for indicated time
periods. Total mRNA was isolated and IFN-γ mRNA expression determined by
qRT-PCR. Data showed that t
1/2
of IFN-γ mRNA declined from 52 min to 42min in
response to Epo.

My results indicated Epo-induced IFN-γ mRNA expression is under
transcriptional regulation by ATF3 binding to the IFN-γ promoter. This induction
occurred in a time-dependent manner in erythroid cells with maximum expression
observed 1 hr post Epo addition. Next, I determined whether Epo affected the
stability of IFN-γ mRNA. As shown in Figure 16, treatment of K562 cells with Epo
40

for 1 hr resulted in a significant de-stabilization in IFN- γ mRNA with a decrease
in the half life of the mRNA (t
1/2
)

from 52 min to 42 min, The kinetics show a
biphasic decline in IFN-γ mRNA, likely involving miRNA-dependent mechanisms
of post-transcriptional regulation.

41

Chapter 4: Erythropoietin and VEGF-mediated
Expression of Placental Growth Factor and its
Regulation by HIF-1α in Erythroid Cells

A. Hypothesis 2

Studies show that levels of PlGF are high in SCD patients, and that PlGF
contributes to inflammation and pulmonary hypertension both in vitro and in vivo
(Sundaram et al., 2010). However, relatively less is known how PLGF expression
is regulated. As erythropoietin (Semenza et al., 1991) and VEGF (Ng et al.,
2006) levels are also elevated in SCD, due to underlying conditions of hypoxia
and increased erythropoiesis, we hypothesized that these molecules may
contribute to increased expression of PlGF in erythroid cells. Also, the promoter
of PlGF has cis-consensus binding elements for HIF-1α. Hence we hypothesized
whether HIF-1α is involved in the transcriptional regulation of PlGF in erythroid
cells. To address this, I have developed the following specific aim.

B. Specific Aim 2

To determine the effect of erythropoietin and VEGF on expression of PlGF in
erythroid cells. Specifically determine the cell-signaling pathway for the
transcriptional regulation PlGF expression by HIF-1α. Also determine the role of
HRE sites in PlGF promoter activity.

42

C. Results

1. Erythropoietin/VEGF mediated PlGF mRNA expression in K562 erythroid
cell line.

Figure 17: Epo/VEGF-induced expression of PlGF in K562 erythroid cells
(A) and (B) Both Epo and VEGF-mediated PlGF expression were maximal
at 2 hrs. Therefore, this time point is utilized for rest of the experiments.

43

As shown in Figure 17A, Epo treatment of K562 cells showed a time
dependent increase in PlGF expression, which was maximal after 2 hr of
induction. Similarly, VEGF treatment of K562 cells showed maximum expression
of PlGF after 2 hr of treatment (Figure 17B). Subsequent time points (>2 hrs),
showed a gradual decline in PlGF expression (Figure 15A and 15B). Thus all
studies were performed with Epo and VEGF after 2 hr induction.

2. Analysis of the PlGF promoter.

Figure 18: Schematic of PlGF promoter (-2649 to +1 bp)
This figure shows the locations of seven hypoxia response elements
(HRE) in this promoter.

Placental Growth Factor promoter is about 2.6kb long. Promoter was
studied in order to check the HRE sites. To understand the transcriptional
regulation of PlGF by HIF-1α, first 3 HRE sites were looked at. In silico analysis
of the PlGF promoter (-2649 to +1_showed potential transcription factor sites for
MTF-1, NRF-1, ATF3 and HRE (not shown). For the purpose of discussion only
the HRE sites are shown (Figure 18).

44

3. Effect of shRNA for HIF-1α and ATF3 on PlGF mRNA expression.

Figure 19: ATF3 and HIF-1α regulates PlGF expression
(A) K562 cells were transfected with shRNA for ATF3 and HIF-1α followed
by 2 hr Epo induction. Epo-mediated PlGF mRNA expression was attenuated by
shRNAs for ATF3 and HIF-1α whereas the control shRNA shows no effect. (B)
Overexpression of ATF3 (o/e ATF3), utilizing variant 1 and variant 4, did not
significantly augment PlGF mRNA expression. Both ATF3 variants appeared to
act similarly with respect to PlGF induction.

45

Previous studies showed a role of metal transcription factor (MTF-1) in the
transcription of PlGF in trophoblast cells. (Nishimoto et al., 2009). In the present
study, we observed Epo-mediated PlGF expression was reduced to basal level
by shRNA for HIF-1α and its expression was also reduced by shRNA for ATF3
(Figure 19A). However, overexpression of ATF3 plasmids did not increase in
PlGF mRNA expression (Figure 19B). This may be due to the possible binding of
Variant 4 to full-length ATF3, hence counteracting its transcriptional activity, thus
working as a dominant negative. (Hua et al., 2006). Taken together, these data
showed that Epo-mediated PlGF expression was transcriptionally regulated by
HIF-1α and ATF3.

4. Effect of Epo and VEGF on PlGF promoter luciferase activity.

Figure 20: Effect of Epo and VEGF on PlGF transcriptional activity
K562 cells were co-transfected with wt PlGF reporter luciferase plasmid
along with β-galctosidase plasmid, the latter for correcting transfection efficiency.
The luciferase activity was normalized to the promoterless pGL3 vector. Cells
were transfected with wild type PlGF promoter construct (wt, -2650/+1). This was
followed by treatment with Epo/VEGF for 6 hrs. Both luciferase and β-
galctosidase activity were measured as described in ‘Materials and Methods’.
The luciferase activity was normalized to that of the untreated PlGF luc construct.
Data are expressed as Mean ± SEM of n=3.
46

As shown in Figure 18, the wt PlGF promoter has several HRE binding
sites. Transfection of K562 cells with wt PlGF promoter followed by Epo and
VEGF treatment for 6 hrs resulted in a 2 and 1.5 fold increase respectively in
luciferase activity. (Figure 20) Taken together, these results showed that both
Epo and VEGF induced transcriptional activity of the PlGF promoter.

5. Chromatin immunoprecipitation analysis of PLGF with antibody to HIF-
1α.

Chromatin Immunoprecipitation with HIF-1α antibody was carried out to
check the binding efficiency of HIF-1α to PlGF promoter. As shown in (Figure
21A, 21B and 21C), VEGF and Epo both increased HIF binding to the promoter
region of PlGF. All three HRE sites (HRE-1, HRE-2 and HRE-3) showed
increased binding of HIF-1α to the native chromatin in response to VEGF
treatment, while Epo treatment showed increased level of binding only to the
HRE-2 site. Nonetheless the data are consistent with involvement of HIF-1α in
transcription of PlGF.

47

Figure 21: Chromatin Immunoprecipitation analysis showed HIF-1α binding
to the PlGF promoter in the native chromatin
(A), (B) and (C) Chromatin was isolated from Epo and VEGF-treated cells
after 2 hrs of induction. Chromatin samples were immunoprepitated using
antibody to HIF-1α (middle panel) or control rabbit IgG (lower panel). PCR was
performed utilizing primers corresponding to three HRE sites (HRE-1, HRE-2 and
HRE-3). The data shows equal loading of input DNA (upper panel) and minimal
PCR products in IgG immunoprecipitated samples.

6. Chromatin remodeling with HDAC inhibitor TSA.

Tricostatin A (TSA) is an organic compound that serves as an antifungal
antibiotic. It is used to alter gene expression by interfering with the removal of
acetyl groups from histones (histone deacetylases, HDAC) and therefore
alternating the ability of DNA transcription factors to access the DNA molecules
inside chromatin, hence promotes histone acetylation by opening up the
chromatin structure.

We examined whether chromatin remodeling played a role in Epo or
VEGF-mediated PlGF transcription. As shown in Figure 22, TSA alone increased
PlGF expression by >2-fold; however, the levels of PlGF expression were not
48

induced further in the presence of Epo and TSA. (Figure 22A) whereas the level
of PlGF gene expression increased to 3-fold, in response to VEGF+TSA. (Figure
22B) These results showed that chromatin opening was perhaps involved in
PlGF expression, but there was no additive or synergistic effect observed with
Epo. Moreover, the data are inconclusive at the present time and further studies
are warranted to understand the role of chromatin remodeling in transcription of
the PlGF gene.

Figure 22: Chromatin remodeling with HDAC inhibitor, Trichostatin A (TSA)
(A) and (B) TSA, an inhibitor of histone deacetylase, augmented PlGF
expression indicating chromatin remodeling allowed upregulation of PlGF
expression under basal and VEGF treated condition, but no additive or
synergistic effect was observed with Epo.
49

Discussion

Chronic hemolytic anemia and vascular occlusions are hallmarks of sickle
cell disease (SCD). Vaso-occlusion induces tissue hypoxia leading to activation
of endothelium, white blood cells including monocytes, and circulating erythroid
cells (Perelman et al., 2003). Due to hemolysis and tissue hypoxia,
erythropoietin, VEGF and PlGF levels are chronically elevated in SCD, and
exaggerated during painful vaso-occlusive crises. Previous studies indicate that
plasma levels of cytochemokines such as IL-1β, TNF-α, MCP-1, MIP-1β and IFN-
γ, are elevated in SCD compared to matched healthy controls (Selvaraj et al.,
2003). In our laboratory, we showed that erythroid cells produced placental
growth factor, an angiogenic growth factor that activated monocytes from normal
individuals to upregulate the expression of IL-1β, MCP-1, MIP-1β and TNF-α
(Perelman et al., 2003). However, monocytes from SCD patients showed
increased expression of same cytochemokines in the absence of treatment with
PlGF showing that monocytes from SCD are in an activated state (Selvaraj et al.,
2003). Although, it is established that plasma levels of PlGF and IFN-γ are high
in SCD patients, relatively less is known about the molecular mechanism(s) of
PlGF expression and inflammatory cytokine IFN-γ. An understanding of the
molecular mechanism of expression of these important molecules will provide a
rationale therapeutic approach to reduce the expression of these molecules
deleterious in SCD patients.

50

In the present study, we examined the molecular mechanisms of
increased expression of IFN-γ and PlGF in model systems as a means of gaining
a better understanding of physiological sequelae in SCD patients. In our
experiments, we showed that Epo and VEGF induce expression of IFN-γ.
We showed that both Epo and VEGF induced expression of IFN-γ in
human erythroid cell line K562. Although we used K562 cells for ease of culture,
the results that were obtained will need to be validated in primary erythroid cells
from normal and SCD subjects. The expression of IFN-γ in response to Epo was
maximal after 1 hr of induction, while the response to VEGF was maximal after 2
hrs. Next, we examined the mechanism by which Epo and VEGF signaling led to
increased expression of IFN-γ. Bioinformatics analysis of the IFN-γ promoter
(approximately 2 kb) showed the presence of cis-binding sequence motifs
corresponding to ATF3, Jun B, CREB, AP-1 in addition to several hypoxia
response elements (HRE). We utilized shRNAs to ATF3 and HIF-1α, and
showed that HIF-1α shRNA attenuated the transcription of IFN-γ mRNA. On the
other hand, transfection with ATF3 shRNA in erythroid cells augmented IFN-γ
expression. Taken together these data showed that HRE elements and ATF3 cis-
binding elements in the promoter played a role in transcription of IFN-γ. These
studies inferred that ATF3 might act as a repressor or a negative regulator for
induction of IFN-γ in erythroid cells, in response to erythropoietin and VEGF.
These results are contrary to recent studies wherein it was shown that during
Th1 differentiation, mediated by either IL-12 or IFN-α, the expression of ATF3
51

increases which leads to increased expression of IFN-γ, showing that ATF3
acted as a positive regulator of human IFN-γ gene expression (Filen et al., 2010).
By contrast, in another study, ATF3 null NK cells showed increased transcription
and secretion of IFN-γ, which was associated with resistance to murine
cytomegalovirus (MCMV) infection in NK cells, indicating that ATF3 acts as
negative regulator of IFN-γ gene expression (Rosenberger et al., 2008). Thus
ATF3-mediated expression of IFN-γ is likely to be cell context and ligand
dependent.

In my present work, the role of ATF3 in the regulation of IFN-γ expression
in erythroid cells was examined. Transfection of erythroid cells with
overexpression plasmid for ATF3 attenuated Epo-mediated IFN-γ expression,
thus validating our results with an alternative approach that ATF3 acted as a
negative regulator of IFN-γ gene expression. Moreover, we utilized IFN-γ
promoter luciferase reporter assay, covering the promoter region from -1147 to
+38 (a fragment of WT IFN-γ promoter). This region of the 5’-flanking DNA
contains an ATF3 site from -62 to -52, a Jun B site from -200 to -191, an HRE
site from -226 to -222, a CREB site from -256 to -249, and two AP-1 binding sites
at -320 to -311 and -978 to -968. The full length IFN-γ promoter (-1147 to +38)
showed 2-3 fold increases in reporter luciferase activity in response to Epo and
VEGF. The functionality of these transcription factor-binding sites in the reporter
construct was ascertained by generating deletion mutation constructs. The
52

mutant promoters had deletions in four sites and intact putative binding site. For
example, deletion mutation construct with ATF3 had mutation in all other sites
except ATF3. Our studies showed that Epo and VEGF-mediated transcriptional
activation of the IFN-γ promoter required ATF3, Jun B and AP-1. These studies
thus supported a potential role for ATF3 in IFN-γ expression.
We utilized chromatin immunoprecipitation (ChIP) analysis to determine
the in vivo role of ATF3 in the transcription of IFN-γ. Our results showed that
ATF3 was recruited to the cis-binding elements of the IFN-γ promoter in
chromatin obtained from Epo- and VEGF-treated K562 cells. The binding of
ATF3 to the promoter region was observed with a primer pair that amplified a ~
200 bp fragment within the proximal 2 kb of the promoter (-2126 to +250). These
results established that the reduced binding of ATF3 to the TSS-proximal region
of the IFN-γ promoter correlated with increased expression of IFN-γ, in response
to Epo and VEGF. In tandem with the mRNA data, it may be inferred that ATF3
has a role as a negative regulator of IFN-γ gene expression.

Next, we determined that both Epo and VEGF effected a 3-4 fold increase
in expression of PlGF from K562 cells. In silico analysis of the promoter region
of PlGF showed the presence of MTF-1, HRE, and ATF3 sites. Previous studies
have shown a role for MTF-1 in the regulation of PlGF expression in trophoblast-
derived cells (Nishimoto et al., 2009). We determined that Epo-mediated PlGF
expression in K562 cells was reduced with HIF-1α shRNA and ATF3 shRNA.
53

Thus indicating that both transcription factors may act as possible activators of
PlGF gene expression. We further validated these results by overexpressing
ATF3, which reduced PlGF gene expression, confirming our above result.
Since there are seven HRE sites in the promoter of PlGF (-2649 to +1),
we examined the role of these elements in the induction of PlGF. To address
this, a promoter reporter luciferase construct was generated by my laboratory
colleague, Dr. Caryn Gonsalves. This construct, namely pGL3-PLGF-luciferase
was transfected in erythroid cells and 6 hrs post-transfection, cells were treated
with Epo. We observed a 2-3 fold increase in reporter luciferase activity,
indicating Epo-treatment induced the PlGF promoter. In future studies, the roles
of individual HRE sites in the PlGF promoter will be investigated utilizing HIF-1α
shRNA, and by sequential mutation of each individual HRE site in the promoter.
Additionally, I utilized the ChiP method to determine the role of HIF-1α binding to
HRE binding sites in the PlGF promoter. Of the three HRE sites that were
examined we observed increased binding of HIF-1α to these HRE sites after cells
were treated with Epo/VEGF. Thus our data showed that Epo and VEGF-
mediated PlGF gene expression in erythroid cells likely required HIF-1α for
transcription of this gene. These studies for the first time, to the best of my
knowledge, showed that HIF-1α was involved in Epo and VEGF-mediated PlGF
expression. Further studies are ongoing to substantiate these results.

54

In conclusion, our studies showed that erythropoietin and VEGF-mediated
IFN-γ expression is transcriptionally regulated by both HIF-1α and ATF3 in K562
erythroid cells. Several experimental approaches were employed to demonstate
that ATF3 acted as a negative regulator of IFN-γ gene expression in erythroid
cells. Further studies are warranted to determine how ATF3 acts as a repressor
of the IFN-γ gene. It is possible that chromatin remodeling or post-translational
modification of ATF3, i.e., phosphorylation, may affect recruitment of transcription
factors such as CREB/JunB to the promoter of IFN-γ, subsequently affecting the
transcription of IFN-γ. An understanding of such pathways will provide new
therapeutic avenues to reduce inflammation in SCD patients.

55

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Abstract (if available)

Abstract Sickle cell disease (SCD) is a genetic disorder, characterized by a mutation within the β-globin chain of the hemoglobin molecule. The clinical manifestations of SCD include hemolytic anemia, vaso-occlusive crises and pulmonary hypertension (PHT). To date, there is no permanent treatment for the disease. SCD patients develop inflammation, PHT and reactive airway disease. Our laboratory findings have shown that levels of placenta growth factor (PlGF) are elevated in SCD, and mediate its effect on endothelial cells and monocytes, in the expression of cytochemokines, endothelin-1 and PAI-1 by activating HIF-1α, independent of hypoxia. Moreover, it has been shown that plasma levels of interferon-γ (IFN-γ) among other cytokines are increased. However, relatively less is known how expression of PlGF and IFN-γ are regulated, as understanding of cell signaling pathways may provide insight to reduce the expression of these molecules, which contribute to inflammation and PHT in SCD. We examined the effect of erythropoietin (Epo) and VEGF, molecules whose expression increases during hypoxia, seen in SCD. ❧ In the present work, we showed that Epo and VEGF upregulated the expression of IFN-γ in K562 erythroid cell line. Furthermore, shRNA for HIF-1α reduced IFN-γ expression while ATF3 shRNA augmented IFN-γ expression. Overexpression of ATF3 attenuated IFN-γ expression. Promoter analysis of IFN-γ utilizing IFN-γ promoter luciferase plasmid, and ChIP, showed occupancy of ATF3 in the promoter region of IFN-γ. Our studies for the first time, to the best of our knowledge, showed that both HIF-1α and ATF3 were involved in the IFN-γ expression, where ATF3 acted as a negative regulator of IFN-γ expression in erythroid cells. ❧ Next, we examined the mechanism of Epo and VEGF-mediated PlGF expression in erythroid cells. Our studies indicated that both Epo and VEGF up regulated the expression of PlGF in K562 cells. Moreover, we inferred that both HIF-1α and ATF3 were involved in its expression, as shRNA for HIF-1α and ATF3 reduced PlGF expression. In silico analysis of PlGF promoter revealed the presence of seven HRE sites in its promoter (~2 Kb). Promoter analysis utilizing wild type, full-length promoter luciferase construct and mutation of selected HRE sites in the promoter further confirmed the role of HIF-1α in PlGF expression. Chromatin immunoprecipitation analysis (ChIP) utilizing antibody to HIF-1α established that HIF-1α was bound to the PlGF promoter in native chromatin to up-regulate the expression of PlGF. ❧ In conclusion, these results showed that Epo and VEGF mediated expression of IFN-γ and PlGF in erythroid cells was transcriptionally regulated by HIF-1α and ATF3. Interestingly, ATF3 acted as a repressor or negative regulator of IFN-γ expression in erythroid cells, mediated by Epo and VEGF.

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Asset Metadata

Creator Jaisinghani, Ruchika (author)

Core Title Transcriptional regulation of IFN-γ and PlGF in response to Epo and VEGF in erythroid cells

School Keck School of Medicine

Degree Master of Science

Degree Program Molecular Microbiology and Immunology

Publication Date 05/03/2014

Defense Date 03/15/2013

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

Tag biochemistry,Biology,cell biology,cell culture,erythroid cells,IFN gamma,microbiology,Molecular Biology,molecular cloning,OAI-PMH Harvest,PlGF,sickle cell

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Tahara, Stanley M. (committee chair), Kalra, Vijay K. (committee member), Machida, Keigo (committee member)

Creator Email rjaising@usc.edu,ruchika.jaisinghani18@gmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c3-250117

Unique identifier UC11288156

Identifier etd-Jaisinghan-1649.pdf (filename),usctheses-c3-250117 (legacy record id)

Legacy Identifier etd-Jaisinghan-1649.pdf

Dmrecord 250117

Document Type Thesis

Rights Jaisinghani, Ruchika

Type texts

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

Access Conditions The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the a...

Repository Name University of Southern California Digital Library

Repository Location USC Digital Library, University of Southern California, University Park Campus MC 2810, 3434 South Grand Avenue, 2nd Floor, Los Angeles, California 90089-2810, USA