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The role of miRNA and its regulation in pulmonary hypertension in sickle cell disease

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The role of miRNA and its regulation in pulmonary hypertension in sickle cell disease

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THE ROLE OF MIRNA AND ITS REGULATION IN PULMONARY
HYPERTENSION IN SICKLE CELL DISEASE

by

Chen Li

____________________________________________________________________

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
(GENETICS, MOLECULAR, AND CELLULAR BIOLOGY)

December 2015

Copyright 2015 Chen Li

ii

ACKNOWLEDGEMENTS
I would like to express my special thanks to Dr. Vijay Kalra, my Ph.D. mentor, whose
erudite knowledge and warm-hearted care in both academic and personal life within the past
several years made my scientific endeavor in the United States and at the University of Southern
California an extremely enjoyable journey and full of excitement. I would also like to thank Dr.
Stanley Tahara. His instructive and rigorous requirements towards scientific research always
reminded me to be a serious researcher along this journey, with his guidance I can always find
my insufficiencies in scientific research. I shared a lot of joy and hard work with them, and
gained constructive advice and encouragement from these two professors. Thus, I owe my
deepest gratitude to them. I want to thank Dr. Wange Lu for serving on my committee and kindly
providing me a lot of valuable insights, and help towards my project design and dissertation
preparation.
I am thankful to my lab members in the past years, Caryn Gonsalves, Jay Hu and Susan
Zhang who helped me in this project, as well as all the professors at the Keck School of
Medicine for their contribution to my education and passing on their knowledge to me, which
profoundly contributed to broadening my horizons in my area of study. Finally, I am eternally
grateful to my parents, Minqiang Li and Jing Cai, who supported me unconditionally in my
education and research, which tremendously motivated me to push forward and confront all
challenges confidently in this journey.

iii

Table of Contents
Acknowledgements ii
List of Figures v
List of Tables vii
List of Abbreviations viii
Abstract xi
Chapter 1: Background and Overview 1
1.1 Sickle Cell Disease 1
1.2 Pulmonary Hypertension 3
1.3 Placenta Growth Factor (PlGF) 4
1.4 Endothelin-1 5
1.5 The PlGF—HIF-1α—ET-1/PAI-1 axis in SCD 7
1.6 miRNAs: biogenesis and potential therapeutic approaches 9
1.7 miRNA-648 and miRNA-199a 12
1.8 Peroxisome Proliferator Activator Receptor α 13
1.9 ATF3 and HDACs 15

Chapter 2: Hypothesis and Specific Aims
2.1 Hypothesis 18
2.2 Specific Aims 18

Chapter 3: Post-Transcriptional Regulation of Endothelin-1 by Placenta Growth Factor
via miR-648 and its transcriptional co-regulation with MICAL3 by PAX5 19
3.1 Introduction 19
3.2 Material and Methods 22
3.3 Results 30
iv

3.4 Discussion 52

Chapter 4: Peroxisome Proliferator-activated Receptor- α –mediated Transcription
of miR-199a2 Attenuates Endothelin-1Expression via Hypoxia-inducible Factor-1α 56
4.1 Introduction 56
4.2 Material and Methods 59
4.3 Results 67
4.4 Discussion 94

Chapter 5: Placenta growth factor mediated transcription of DNM3os/miR-199a2
is negatively regulated by ATF3 and associated histone deacetylase 6
in sickle cell disease 100
5.1 Introduction 100
5.2 Material and Methods 102
5.3 Results 109
5.4 Discussion 132

Chapter 6: Summary of Study 137

Reference 140
v

List of Figures
Figure 1: The important clinical consequences of Sickle Cell Disease 1
Figure 2: The PlGF-induction of cytochemokines expression in monocytes 8
Figure 3: The biogenesis of miRNA 11
Figure 4: PlGF-mediated expression of candidate miRNAs in human endothelial
cell line (HMEC-1) with predicted binding site(s) within the 3’UTR of ET-1 mRNA 31
Figure 5: Effect of exogenous miR-648 on ET-1 in HMEC-1. 34
Figure 6: Endogenous miR-648 expression and effects of exogenous miR-648 and
anti-miR-648 on ET-1 mRNA and protein in cultured primary human endothelial cells 38
Figure 7: Effects of miR-648 and anti-miR-648 oligonucleotides on ET-1-3’UTR
reporter luciferase activity 40
Figure 8: Role of P1, P2 and internal P2a promoter in transcription of MICAL3 42
Figure 9: Expression of PAX5 in endothelial cells 45
Figure 10: PAX5 regulates transcription from MICAL3-P1 promoter and expression
of pre-miR-648 48
Figure 11: miR-648 plasma levels in SCA and control subjects, and working model
of miR-648 mediated regulation of ET-1 expression 51
Figure 12: PlGF attenuates miR-199a2-5p expression, which targets the 3’UTR
of HIF-1α mRNA 68
Figure 13: miR-199a2 targets the 3’UTR of HIF-1α mRNA affecting expression
of downstream target gene ET-1 71
Figure 14: PlGF attenuates expression of miR-199a2 and its host gene DNM3os
in endothelial cells 75
Figure 15: PPARα regulates the transcription of miR-199a2 and its host gene DNM3os 79
Figure 16: PPARα cis-binding elements in promoter ofDNM3osregulate expression
of DNM3os as determined by reporter assay and ChIP 82
Figure 17: PPARα agonists attenuate expression of HIF-1α and ET-1 in both HMEC
vi

cells and primary HLMVEC 86
Figure 18: miR-199a, HIF-1α, and ET-1 biomarkers in human SCD subjects
and fenofibrate-treated sickle mice 92
Figure 19: PlGF mediated expression of ATF3 involves signaling via PI-3-Kinase,
MAP kinase, p38MAPK and Jun-terminal kinase-1 110
Figure 20: ATF3 acts as repressor of DNM3os and miR-199a2 transcription 113
Figure 21: ATF3 transcription factor binding to cis-binding elements in the promoter of
DNM3os 116
Figure 22: Association of histone deacetylase in chromatin remodeling and
transcription of DNM3os/miR-199a2 119
Figure 23: Effect of histone deacetylase inhibitors on DNM3os transcriptional activity 121
Figure 24: Binding of transcription factors, ATF3, JDP2, c-Jun, HDAC6 ATF2
and HDAC7 to DNM3os promoter in native chromatin as assessed by ChIP 124
Figure 25: Effect of Tubacin, ATF3 shRNA and HDAC6 shRNA on chromatin
structure of ATF3 binding site 1 and 2 126
Figure 26: qRT-PCR of HIF-1α and ET-1 mRNA in PlGF treated HMEC and
with transfection of ATF3 shRNA 130

vii

List of Tables
Table 1: Oligonuleotide primers used in the Chapter 3 study 24
Table 2: Oligonuleotide primers used in the Chapter 4 study 63
Table 3: Oligonucleotide primers used in the Chapter 5 study 104
Table 4: Proteomic analysis of HMEC cells transfected with ATF3 expression
plasmid, followed by immunoprecipitation with ATF3 antibody and mass
spectroscopic analysis 119
viii

List of Abbreviations
3' UTR 3' untranslated region
5’ UTR 5’ untranslated region
ACS acute chest syndrome
AP-1 activator protein-1
ATP adenosine triphosphate
ATF3 activating transcription factor 3
C/EBP CCAAT box-enhancer-binding protein
CBP CREB- binding protein
ChIP chromatin immunoprecipitation assay
DNM dynamin
EGTA ethylene glycol tetra acetic acid
Epo erythropoietin
ET-1 endothelin-1
ETB receptor endothelin B receptor
FAIRE formaldehyde assisted isolation of regulatory elements
GAPDH glyceraldehyde 3-phosphate dehydrogenase
HbAS sickle cell trait
HbSS sickle hemoglobin
HDAC histone deacetylase
HIF hypoxia inducible factor
HLH helix-loop-helix
HPMVEC human pulmonary microvascular endothelial cells
ix

HRE hypoxia response element
HSCT hematopoietic stem cell transplantation
HUVEC human umbilical vascular endothelial cell
ICAM-1 inter-cellular adhesion molecule-1
IL interleukin
JNK c-jun N-terminal kinase
MCP-1 monocyte chemotactic protein -1
MICAL3 Microtubule-associated monoxygenase, calponin and LIM domain
containing 3
MIP-1β macrophage inflammatory protein-1β
miRNA microRNA
NADPH nicotinamide adenine dinucleotide phosphate
NF-κB nuclear factor κB
NO nitric oxide
PAP pulmonary artery pressure
PHD prolyl hydroxylase
PHT pulmonary hypertension
PI3K phosphoinositide 3-kinase
PlGF placenta growth factor
PPAR peroxisome proliferation activator receptor
pre-miR precursor microRNA
pri-miR primary microRNA
qRT-PCR quantitative real time polymerase chain reaction
x

RBCs red blood cells
RISC RNA-induced silencing complex
SCD sickle cell disease
siRNA small interfering RNA
SSRBCs sickle red blood cells
VEGF vascular endothelial growth factor
VHL von Hippel Lindau protein
WBCs white blood cells
xi

ABSTRACT
Pulmonary hypertension (PHT) is a highly prevalent complication of Sickle Cell Disease
(SCD), and it is a major cause of early morbidity and mortality in sickle cell patients. SCD
patients with PHT show increased levels of Placenta Growth Factor (PlGF) and a potent vaso-
constrictor Endothelin-1 (ET-1) in the plasma. The previous work from our laboratory has shown
that PlGF, elaborated from erythroid cells, show high levels in the plasma of SCD patients
compared to healthy subjects. PlGF has been shown to induce the expression of inflammatory
cytochemokines and ET-1 through a mechanism involving the activation of hypoxia induced
factor-1α (HIF-1α), independent of hypoxia. Moreover, studies showed that binding of HIF-1α to
the hypoxia response elements (HRE) in the promoter region of the ET-1 gene leads to increased
synthesis of ET-1 mRNA.
However, in the PlGF—HIF-1α—ET-1 axis of PHT, the mechanisms by which PlGF
enhances the stability of ET-1 and HIF-1α mRNA, and how HIF-1α regulates its down-stream
effector ET-1 remain unclear. In Chapter 3, we showed that in human endothelial cells, the
stability of ET-1 mRNA was increased in response to PlGF treatment via regulation of miR-648,
which decreases ET-1 expression by direct binding to the 3’ untranslated region (3’UTR) of ET-
1 mRNA. Moreover, our data showed that in endothelial cells, miR-648 is co-transcribed with
Microtubule-associated monooxygenase, calponin and LIM domain containing 3 (MICAL3);
expression of miR-648 and MICAL3 are both repressed by PlGF in endothelial cells, and a
transcription factor (TF); Pair Box 5 (PAX5) is involved in the regulation of this process. Further,
we extend the study to SCD patients, and concluded that SCD patients have a significantly
higher level of miR-648 in plasma compared to healthy subjects. Thus, miR-648 can be a
xii

promising biomarker of SCD and targeting the transcription of miR-648 or its host gene
MICAL3 may provide novel insights into new therapeutic approaches for treatment of SCD.
Our group and others report miR-199a is involved in regulation of Hypoxia Inducible
Factor-1α (HIF-1α) in alcoholic liver disease and tumorigenesis. HIF-1α was found to be the
upstream regulator of ET-1 in response PlGF induction of ET-1 in endothelial cells, and it is
induced by PlGF independently of hypoxia. In Chapter 4, we describe that in endothelial cells,
PlGF reduces the endogenous level of miR-199a, which targets HIF-1α mRNA by direct binding
to its 3’UTR and concomitantly regulates expression of the downstream effector ET-1.
Moreover, our data indicated that miR-199a2 is co-transcribed with its host gene Dynamin 3
opposite strand (DNM3os) and regulated by the transcription factor, peroxisome proliferator-
activated receptor α (PPARα). Additionally, we show that fenofibrate, a PPARα agonist,
increased the expression of miR-199a2 and DNM3os; the former was responsible for reduced
expression of HIF-1α and ET-1. In vivo studies of fenofibrate-fed Berkeley sickle mice resulted
in increased levels of miR-199a2 and reduced levels of ET-1 in lung tissues. This study
provided a potential therapeutic approach whereby fenofibrate-induced miR-199a2 expression
can ameliorate PH by reduction of ET-1 levels.
Since PlGF repression of miR-199a and its host gene DNM3os is a key process in PlGF
induction of HIF-1α and ET-1 in SCD PHT, it is necessary to understand how this process is
regulated. PlGF strongly induced expression of the transcriptional repressor activating protein
(ATF3) in endothelial cells. Proteomic analysis of cell lysate derived from HMEC cells
expressing ATF3 and co-immunoprecipitated with ATF3 antibody, showed association of ATF2,
JDP2, HDAC6 and HDAC7 with ATF3. Furthermore, we showed histone deacetylase (HDAC),
specifically HDAC6, was associated with the ATF3 site in the DNM3os promoter to repress
xiii

DNM3os transcription. Tubacin, a selective inhibitor of HDAC6, induced the expression of
DNM3os RNA and premiR-199a2 RNA, and subsequently reduced the PlGF-mediated
expression of HIF-1α and its target gene ET-1. In vivo studies of lung tissue from Berkeley
Sickle (SS) mice showed increased levels of ATF3, which correlated with increased expression
of ET-1, a marker of pulmonary hypertension. Our studies provided a potential therapeutic
approach whereby Tubacin-induced miR-199a2 expression thereby attenuating endothelin-1
levels, a potent vasoconstrictor, in SCD.
In conclusion, our studies identify miRNAs that regulate the expression of HIF-1α and
ET-1. Furthermore, these studies for the first time demonstrate the transcriptional regulation of
miRNAs by specific transcription factors e.g. PAX-5 and PPARα and histone deacetylase
(HDACs). These studies provide a new therapeutic paradigm to upregulate the expression of
these miRNAs and concomitantly attenuating the expression of HIF-1α and its downstream
target gene e.g. ET-1, the latter involved in pulmonary hypertension in sickle cell disease.
1

Chapter 1 Background and Overview

1.1. Sickle Cell Disease
Sickle cell disease (SCD) is a genetic disorder caused by a mutation in the β-globin chain
of the hemoglobin molecule, wherein the sixth amino acid residue, glutamic acid is replaced by
valine. The abnormal hemoglobin (HbS) polymerizes at low oxygen tension forming a 14-
stranded fiber that distorts the shape of the red blood cells giving rise to a sickle shape. When the
deformed sickle red blood cells pass through small capillaries, they undergo hemolysis leading to
hemolytic anemia. Furthermore, adhesion and trapping of these SS RBCs in the small capillaries
and venules leads to localized ischemia or hypoxia, which further induces sickling, and thus a
Figure 1. Important clinical consequence of SCD includes vaso-occlusion, stoke, acute chest
symptom, infection, pulmonary hypertension, etc. Adapted from Makani et. al. The Scientific World
Journal (2013)
Sickle red blood cell
Normal red blood cell
2

vicious cycle develops (Francis & Johnson, 1991; Platt et al., 1991; Wong et al., 1992). At
present, the only cure for SCD is hematopoietic stem cell transplantation (HSCT) or gene
therapy utilizing transduced autologous hematopoietic stem cells. However, since the gene
therapy approach is still at a very early stage of development and HSCT is greatly limited by the
availability of a genetically matched donor, there is still a long way for developing a complete
cure of SCD (Shenoy, 2013).
Vascular occlusion is caused by the blockage of the microvasculature by sickle
erythrocytes and triggers a series of severe complications such as painful vaso-occlusive crises,
acute chest syndrome and irreversible organ damage, leading to stroke and renal failure. In the
US, 8% of Afro-Americans are carriers for HbS, while 100,000 individuals are homozygous for
SCD. The median life span of homozygous SCD individuals in the US is approximately 50
years. However, in sub-Saharan Africa 180,000 infants are born per year with SCD, and 50% of
them die before 5 years of age (Modell & Darlison, 2008).
Hydroxyurea, which causes an increase in hemoglobin F (HbF) levels in SS RBCs, has
been approved for use in humans. The increased amount of HbF in SS RBC delays
polymerization of HbSS. However, this drug is effective in only 50% of SCD patients.
Moreover, this drug has been found to be a teratogenic agent in animal studies. Thus
hydroxyurea has the potential for side effects in SCD patients (Steinberg, Nagel, & Brugnara,
1997). For these reasons, new drugs should be developed for treatment of SCD. In order to
determine the targets of new drugs beneficial for treatment of SCD, we have to first understand
the contribution of SS RBC’s biology to vascular dysfunction and pathophysiology of SCD.

3

1.2. Pulmonary Hypertension
Pulmonary Hypertension (PHT) is a progressive condition. PHT is defined, as below
normal or reduced cardiac output, without any significant left heart disease, lung disease, or
chronic thromboembolic disease, with the mean pulmonary artery pressure (mPAP) no greater
than 25 mm Hg (Chin & Rubin, 2008; McLaughlin & McGoon, 2006). In 2003, the Third World
Symposium on Pulmonary Arterial Hypertension held in Venice, Italy categorized PHT into five
different groups: (1) Pulmonary arterial hypertension (PAH); (2) Pulmonary hypertension with
left heart disease; (3) Pulmonary hypertension associated with lung disease and/or hypoxemia; (4)
Pulmonary hypertension due to chronic thrombotic and/or embolic disease; and (5)
Miscellaneous. Further classification of PHT subgroups is based on its causes, for example,
idiopathic pulmonary arterial hypertension (IPAH), familial pulmonary arterial hypertension
(FPAH), pulmonary arterial hypertension associated with HIV, drugs and toxins, chronic
obstructive lung disease, and interstitial lung disease. (Simonneau et al., 2004). Common
symptoms of PAH include dyspnea, fatigue, non-productive cough, angina, and peripheral
edema (McLaughlin & McGoon, 2006).
PHT is a highly prevalent complication of SCD, and it is a major cause of morbidity and
mortality in sickle cell patients. It occurs in approximately 10% of adults patients with SCD, and
its diagnosis is associated with a 38-40% mortality in the following 2-6 years after PHT
diagnosis (Gladwin et al., 2004; Minniti et al., 2009). Studies also showed sickle mice develop
PHT with increasing age, manifested as high pulmonary artery pressures and right ventricular
hypertrophy (Sundaram et al., 2010). Physiological factors implicated in PHT in SCA include
endothelial dysfunction, pulmonary vasoconstriction, and vascular remodeling; all of which are
4

associated with chronic hemolysis, hypoxia, hemostatic activation, and inflammation (Ataga et
al., 2008; Castro & Gladwin, 2005; Hsu et al., 2007; Minniti et al., 2009).
1.3. Placenta Growth Factor (PlGF)
Placenta growth factor (PlGF), an angiogenic growth factor, is a member of platelet
derived growth factor/vascular endothelial growth factor (PDGF/VEGF) family. PlGF has a high
sequence similarity to VEGF; it binds only to vascular endothelial growth factor receptor-1
(VEGFR-1), while VEGF binds to both VEGFR-1 and VEGFR-2, with a higher affinity for
VEGFR-2 over VEGFR-1. Furthermore, PlGF has been shown to cause increased secretion of
VEGF from monocytes. PlGF is highly expressed in placental trophoblasts and human umbilical
vein endothelial cells and in non-placental tissues such as mouse embryonic fibroblasts and
erythroblasts (De Falco, 2012; Green et al., 2001; Tordjman et al., 2001). PlGF expression is
induced by hypoxia, erythropoietin and iron (Kelly et al., 2003; Perelman et al., 2003; X. Wang
et al., 2014; Yamakawa et al., 2003). PlGF induces cell signaling via binding to its cognate
receptor VEGFR-1 (De Falco, 2012; Park, Chen, Winer, Houck, & Ferrara, 1994).
Recent studies have shown plasma levels of PlGF are abnormally high in patients with
hemolytic anemia such as sickle cell disease due to increased erythroid cell hyperplasia
(Perelman et al., 2003). Our lab has previously shown increased PlGF levels in the bone marrow
from SCD subjects. Moreover, PlGF activates peripheral blood nucleated cells, specifically
monocytes, and results in increased expression of pro-inflammatory cytokines and chemokines
by these cells (Perelman et al., 2003; Selvaraj et al., 2003). Furthermore studies show high
plasma PlGF levels correlate with increased incidence of vaso-occlusive events in SCD
subjects(Perelman et al., 2003).
5

Studies from our group on humanized sickle (Berkeley-SS) mice showed plasma levels of
PlGF and ET-1 were elevated significantly compared to healthy controls. The specific
contribution of elevated PlGF to pulmonary hypertension has been further validated by
stimulating erythroid expression of PlGF using a lentiviral vector in normal mice to the abnormal
levels observed of PlGF in Berkeley-SS mice. Normal mice over-expressing PlGF with this
lentiviral vector also showed increased production of ET-1 and developed increased right
ventricular (RV) pressures, RV hypertrophy and pulmonary fibrosis, which is consistent with the
symptoms of pulmonary hypertension in SCD (Sundaram et al., 2010). Additionally, studies
conducted by another research group also found a positive association between high PlGF levels
to pulmonary hypertension in SCA patients (Brittain, et al., 2010), thus strengthening the
observations reported from our group. These studies strongly support the role of PlGF-induced
ET-1 in pulmonary hypertension in sickle animal models and human patients. However,
additional studies are needed to understand the molecular mechanisms of ET-1 expression
regulated by PlGF, which will also provide novel insights in repressing the development of PHT
in SCD.

1.4. Endothelin-1
Endothelin-1 (ET-1) is processed into the active form from preproendothelin-1 (PPET-1)
and encoded by the EDN1 gene in human. There are three isoforms of endothelin peptides: ET-1,
ET-2, and ET-3, and two endothelin receptors (ET-AR and ET-BR) that are present in human. It
has been shown that ET-1 and ET-2 are 90% homologous in amino acid sequence, and ET-3 is
70% identical to ET-1 and ET-2 (Shao, Park, & Wort, 2011). ET-1 is primarily produced by
vascular endothelial cells and smooth muscle cells, airway epithelial cells and smooth muscle
6

cells, macrophages, and fibroblasts (Shao et al., 2011). ET-1 was identified in 1988 by Dr.
Yanagisawa's group as a 24kD endothelium-derived secreted peptide, which is the predominant
isoform among the three and is the most potent vasoconstrictor known to date (Yanagisawa et al.,
1988).
Studies show that there is a significant increase in plasma levels of ET-1 in patients with
hypertension, atherosclerosis, heart disease, and SCD (Barton & Yanagisawa, 2008). ET-1 has
also been identified to play a role in cell growth and invasion in many tumors, including ovarian,
prostate and breast cancer (Bagnato & Rosanò , 2008). Previous studies concluded that a number
of factors and conditions can regulate the cellular level of ET-1, such as transforming growth
factor- β (TGF-β) (Castañ ares et al., 2007), tumor necrosis factor-α (TNF-α) and interleukin-8
(IL-8) (R.-z. Zhao, Chen, Yao, & Chen, 2005), insulin (Nagai et al., 2003) and hypoxia (Aversa
et al., 1997). Moreover, it has been shown that cis-regulatory elements contribute to the
regulation of ET-1 by direct binding to specific promoter regions located upstream of the
transcription start site (TSS), such as activator protein-1 (AP-1), nuclear factor-1 (NF-1) and
GATA binding protein-2 (GATA-2) (M. E. Lee, Bloch, Clifford, & Quertermous, 1990).
Hypoxia is one of the more potent inducers of ET-1; its expression has been shown to be
elevated by hypoxia inducible factor-1 α (HIF-1α) via binding to the Hypoxia Response
Elements (HRE) to its promoter (Hu, Discher, Bishopric, & Webster, 1998; Yamashita, Discher,
Hu, Bishopric, & Webster, 2001). These studies showed there is binding of HIF-1α at a HRE
positioned at -118 to -125 bp upstream from the 5’ transcription start site (TSS) of ET-1 gene, in
response to hypoxia. Moreover, previous studies from our laboratory also showed a significant,
high level of ET-1 in the plasma of patients with SCD, which correlates with a high level of
7

pulmonary artery pressure (PAP), an indicator of pulmonary hypertension (Patel, Gonsalves,
Malik, & Kalra, 2008).
The important role of ET-1 in PHT was also supported by studies showing endothelin-1
receptor antagonists are beneficial in the treatment of primary pulmonary hypertension (Benza et
al., 2008) and endothelin-1 receptor antagonists have also been found to be effective in sickle
Antilles-hemoglobin-D mice exhibiting PHT (Sabaa et al., 2008). Thus, by lowering ET-1 or its
upstream activator HIF-1α in PHT of SCD patients, it is expected that this novel approach will
repress or delay the development or progression of PHT.

1.5 The PlGF—HIF-1α—ET-1/PAI-1 axis in SCD
Previous studies from our group showed that normal mice overexpressing PlGF by
lentiviral vector show an upregulation of their plasma ET-1 levels (Sundaram et al., 2010).
Consistent with these findings, plasma levels of PAI-1 and ET-1 are high in a mouse model of
sickle cell disease, i.e. Berkeley Sickle mice (BK-SS), which also shows higher plasma PlGF
levels (N. Patel et al., 2010; Sundaram et al., 2010). Conversely, PlGF knockout mice, which
have lower plasma PlGF levels, exhibit lower plasma levels of PAI-1 and ET-1 (N. Patel et al.,
2010; Sundaram et al., 2010). Furthermore, we show augmentation of erythroid PlGF expression
by lentivirus vector, in normal mice, raised PlGF to the levels seen in sickle mice, resulting in
increased production of endothelin-1 with associated pulmonary changes as seen in pulmonary
hypertension in SCD (Sundaram et al., 2010). These studies were corroborated in a study with
123 SCD patients, wherein higher plasma PlGF levels were associated with anemia, higher
plasma ET-1 levels, and tricuspid regurgitant velocity, the latter reflective of peak pulmonary
8

artery pressure (Sundaram et al., 2010). Moreover, in order to understand the molecular
mechanism of PlGF induction of its downstream targets, our lab also conducted additional
investigation, and showed PlGF induces ET-1 and plasminogen activator inhibitor-1 (PAI-1) via
the activation of HIF-1α, independently of hypoxia (Patel et al., 2008; N. Patel et al., 2010). ET-
1 released from pulmonary endothelial cells activates monocytes via ET-BR receptor to
upregulate the expression of cytochemokines as depicted in Figure 2.

Figure 2. The PlGF-induction of cytochemokine expression in monocytes. PlGF activates the NAPDH-
oxidase pathway to produce ROS and activates PI-3 kinase. Both of these pathways activate HIF-1 α
which translocate into the nucleus and forms heterodimers with HIF-1β. The complex binds to HRE in
ET-1 promoter. Adapted from Patel et. al. Blood. (112) 856-865. 2008. Gonsalves and Kalra. J
Immunology. (185) 6253-6264.2010

9

These in vitro and in vivo studies support the important role of PlGF in up-regulating the
expression of ET-1 in endothelial cells and associated expression of inflammatory cytokines and
chemokines.

1.6. miRNAs: biogenesis and potential therapeutic approaches
MicroRNAs (miRNAs) are a large family of small RNAs of ~22 nucleotides in length
that play an important role in gene regulation by complementary binding to the target mRNA.
The first miRNA, lin-4 was discovered in 1993 by Ambros and his coworkers (R. C. Lee,
Feinbaum, & Ambros, 1993). Within two decades, numerous miRNAs were discovered to be
involved in crucial cellular pathways, including tumorigeneses/pathogenesis processes through
regulation of levels of key transcription factors, e.g., miR-34 (Misso et al., 2014). Thus,
elucidation of these types of relationships is a promising avenue for development of novel
therapeutic approaches based on miRNAs.
miRNAs are generated from miRNA coding genes, which are scattered within various
genome contexts, including non-coding, intron regions and exon regions, many are also found in
independent clusters and transcription units. RNA polymerase II is the major polymerase
involved in transcription of miRNA coding genes. Transcription of miRNA coding genes results
in synthesis of a primary miRNA (pri-miRNA), which ranges in length from several hundred to
several Kbp (Cai, Hagedorn, & Cullen, 2004). Pol II transcribed pri-miRNAs are characterized
by a 5’ cap structure and 3’ polyadenylated tails (Y. Lee et al., 2004) as shown in Fig. 3. The
pri-miRNA is then processed within the nucleus by a multiprotein complex, Microprocessor. The
central enzymatic component of Microprocessor is Drosha, which has an RNase III-like activity.
10

Other components of Microprocessor include double-stranded RNA-binding domain (dsRBD)
protein GCR8/Pasha (Denli, Tops, Plasterk, Ketting, & Hannon, 2004). Drosha recognizes and
cleaves the stem-loop structure of pri-miRNA and generates a ∼70-nt hairpin precursor miRNA
(pre-miRNA). The cleavage by Drosha of pre-miRNA leads to a 2 nt 3’-overhang of the stem-
loop structure. The 3’overhang serves as a recognition signal for exportin-5 to translocate pre-
miRNA from the nucleus to cytoplasm in a Ran-GTP-dependent manner (Fig. 3). After export
into the cytoplasm, pre-miRNAs are further processed by another RNase III-like enzyme, Dicer,
that trims the ~70 nt pre-miRNAs into the ~22 nt mature miRNAs (Saito, Ishizuka, Siomi, &
Siomi, 2005) (Fig. 3). One strand of mature miRNA is selected and loaded into the RNA-induced
silencing complexes (RISCs), while the other strand is usually degraded (Fig.3). However, in
some cases, both strands of mature miRNA are functional for regulating target gene expression,
for example, miR-199a-5p is reported to regulate HIF-1α mRNA in lungs from patients with
Chronic Obstructive Pulmonary Disease (COPD) (Mizuno et al., 2012) and miR-199a-3p is
reported to regulate caveolin-2 in endothelial cells and breast cancer cell line (Shatseva, Lee,
Deng, & Yang, 2011). miRNAs function in a form of RNA-induced silencing complexes
(RISCs) (Fig. 3) with other components such as Argonaute (AGO) family proteins. RISC with its
associated guide miRNA function by nearly perfect or partial complementary base-pairing to the
target mRNA. Ideally a 2~8 nt seed region with perfect base pairing between the miRNA
sequence and its target occurs (Filipowicz, Bhattacharyya, & Sonenberg, 2008). The mechanisms
by which miRNA regulates its target mRNA can be direct cleavage, translational repression or
deadenylation (Fig. 3). However, only AGO 2 was exclusively found to be able to cleave target
mRNA because of its RNaseH-like P-element induced wimpy testis (PIWI) domain (Filipowicz
11

et al., 2008). Other AGO proteins in RISCs may function by directing RNases to the designated
mRNA.

Figure 3. The biogenesis of miRNA. The mechanism of miRNA mediated gene silencing. miRNAs
can target the 3’ untranslated regions (3’ UTRs) of mRNAs by either perfect pairing or imperfect
nucleotide pairing in miRNAs to target gene. The miRNA mediated gene silencing occur by different
mechanisms, either by cleaving and degradation, or translational repression of target mRNA. Adapted
from Diederichs et al. Nature Cell Biology. (Mar. 2009)

12

1.7. miRNA-648 and miRNA-199a
miR-648 is located in the first intron of its host gene, Microtubule-associated
monooxygenase, calponin and LIM domain containing 3 (MICAL3). MICAL3 is a member of
the MICAL family of flavoprotein monooxygenases and is involved in axon guidance and actin
remodeling (Hung et al., 2010; Kolk & Pasterkamp, 2007). More recently, MICAL3 has been
shown to cooperate with Rab6 and Rab8 in exocytosis events (Grigoriev et al., 2011a). It was
reported in 2012 that miR-648 is present at relatively high levels in in normal human plasma as
detected by a novel RT-PCR assay of microRNAs using s-poly(T), a specific oligo (dT) reverse
transcription primer (K. Kang et al., 2012). By contrast, its levels are down-regulated in ovarian
carcinoma cell lines and tissues (Dahiya et al., 2008). A recent study published in 2015 showed
that miR-648 levels are statistically different among multiple sclerosis (MS) patients, with higher
miR-648 levels observed in relapsing illness patients (Kacperska et al., 2015).
miR-199a is one of the more well studied microRNAs to date. It has physiological
functions in tumorigenesis(both development and metastasis) (Joshi et al., 2014; B.-K. Kim, Kim,
& Yoon, 2015), cell proliferation and survival (Song et al., 2014), monocyte/macrophage
differentiation (Lin et al., 2014) and cardiac hypertrophy (Da Costa Martins & De Windt, 2012).
These varied cellular functions clearly exemplify its potentially broad biological activity.
Interestingly, miR-199a-5p and miR-199a-3p are synthesized from both premir-199A1
and premir-199A2. These isogenes are synthesized from opposite strand transcripts arising from
an intron of DNM2 or DNM3, respectively. The miRNA encoded by miR-199B is similar in
nucleotide sequence to those encoded by miR-199A1 and miR-199A2 isogenes, but has a
different seed sequence. For miR-199a2 stemming from DNM3os, there is a second miRNA in
13

the DNM3os transcription unit, miR-214, which has been extensively studied in embryonic
development in mice (Y.-B. Lee et al., 2009) and human (Duan et al., 2012).

1.8 Peroxisome Proliferator Activator Receptor α
Peroxisome proliferator-activated receptors (PPAR) are a group of nuclear receptor
proteins that function as transcription factors in the regulation of gene expression. PPARs play
essential roles in the regulation of cell differentiation and metabolic processes, especially lipid
and carbohydrate metabolism. PPARs are also reported to be involved in neoplasia,
inflammation and wound healing, development and tumorigenesis processes (Y.-X. Wang, 2010).
There are three types of PPAR presently known: α, β, and γ (the latter includes four
subtypes γ1, γ2, γ3 and γ4). All PPARs are ligand-activated transcription factors, and they can
heterodimerize with the retinoid X receptor (RXR) for recruitment to peroxisome proliferator
hormone response elements (PPREs) on target gene promoters. The binding of ligand to PPARs
leads to a change in the PPAR: ligand conformation, which further results in the release of co-
repressors and recruitment of co-activators, subsequently, regulating target gene expression.
Studies have shown all PPARs share common structural features: they have an amino-terminal
zinc-finger DNA-binding domain (DBD) which is responsible for the recruitment to target DNA
region, and a carboxyl-terminal ligand-binding domain (LBD) which is responsible for forming
the PPAR: ligand complex (Bensinger & Tontonoz, 2008). Moreover, it was reported in 1997 by
Mukherjee and colleagues that RXR agonists can improve the activity of RXR: PPAR γ complex
in the diabetic mouse model and thus shows antidiabetic effects (Mukherjee et al., 1997).
14

PPARα is the first characterized PPAR and is extensively studied because it was found to
be highly involved in liver and skeletal muscle lipid metabolism. It regulates the expression of
genes encoding enzymes and transport proteins for processes such as fatty acid (FA) oxidation
and glucose homeostasis.
PPARα ligands (agonists) can be either synthetic or endogenous. Importantly, FAs and
FA-derived compounds are natural ligands for PPARα. Thus in the case of lipid uptake, these
natural ligands stimulate and activate PPARα by forming the ligand: PPARα complex to induce
the downstream gene expression. The transcription of these downstream genes is increased, and
the oxidation systems, including microsomal omega-oxidation system, mitochondrial and
peroxisomal beta-oxidation are activated and subsequently fatty acid oxidation (Bensinger &
Tontonoz, 2008; Lefebvre, Chinetti, Fruchart, & Staels, 2006).
The endogenous ligands of PPARα include unsaturated fatty acids (e.g. omega 3 fatty
acids) and eicosanoids (e.g. leukotriene B4) (Grygiel-Gorniak, 2014). Omega-3 fatty acids have
anti-inflammatory effects (Lo Verme et al., 2005). The omega 3 fatty acid: docosahexaenoic acid
(DHA) and eicosapentaenoic acid (EPA) are reported to reduce the risk of coronary heart disease,
hypertension, primary heart attack, and rheumatoid arthritis (Sethi et al., 2002). The synthetic
PPARα agonists are fibrates, including fenofibrate, clofibrate and benzofibrate. It was reported
that these fibrates can decrease the level of triglycerides, low density lipoprotein (LDL) and very
low density lipoprotein (VLDL) in serum through an increase in gene expression of those
involved in fatty acid-β-oxidation, and increased levels of high density lipoprotein (HDL)
(Grygiel-Gorniak, 2014). Thus, these fibrate based drugs, were clinically approved, and are still
widely used alone or along with statins to reduce the risk of cardiovascular disease and as
treatments for hypercholesterolemia and hypertriglyceridemia (Yang & Keating, 2009).
15

1.9. ATF3 and HDACs
Activating transcription factor 3 (ATF3) belongs to the mammalian activation
transcription factor (ATF)/cAMP response element-binding (CREB) protein family of
transcription factors, which includes ATF1, ATF2, CREB, CREM, ATF4, ATF5, ATF6, ATF7
and B-ATF Thompson, Xu, & Williams, 2009). It has been determined that the ATF/CREB
family members share the same structural features: an N-terminal DNA binding domain and a C-
terminal basic leucine zipper (bZIP) domain. The DNA binding domain binds to specific
promoter sequences, such as cyclic AMP response element (CRE), with the consensus sequence
TGACGT A/C A/G, while the leucine zipper region is responsible for forming homodimers or
heterodimers with other bZIP-containing proteins such as the AP-1, C/REM, C/EBP, or Maf
families(T. Hai & Hartman, 2001; Thompson, Xu, & Williams, 2009).
Studies of the full-length cDNA of ATF3 revealed that it is a protein of 181 amino acids,
with an approximate molecular weight 22 KD. Like other ATF/CREB family members, ATF3
dimerizes with other factors containing the bZIP motif to form heterodimers, including other
members of the ATF/CREB family, such as ATF2, c-Jun, JunB, JunD (T. Hai, Wolfgang,
Marsee, Allen, & Sivaprasad, 1999) to regulate target gene expression. It is also pertinent to
mention that different studies conclude these heterodimers can behave either as an activator or
repressor, which depends on the promoter context and the specific factor ATF3 heterodimerizes
with (Lu, Wolfgang, & Hai, 2006; Yin et al., 2010). For example, it was reported that ATF3 can
heterodimerize with JDP2 to repress ATF3 expression itself in MEF cells (Darlyuk-Saadon,
Weidenfeld-Baranboim, Yokoyama, Hai, & Aronheim, 2012). Alternatively, ATF3 can be a
transcription activator when it is interacts with c-Jun or JunB, for transcription of heat shock
16

protein 27 (hsp27) in neuronal cells (Nakagomi, Suzuki, Namikawa, Kiryu-Seo, & Kiyama,
2003).
Extensive studies have shown ATF3 is a stress response gene constitutively expressed at
a low level in quiescent cells. However, with external stress signals, such as cellular signals
initiated by cytokines, genotoxic agents, or physiological stresses, ATF3 can be significantly
induced. It has been shown that ATF3 plays an important role in several pathological conditions
such as host-defense immunity, breast cancer metastasis and hepatic gluconeogenesis (Tsonwin
Hai, Wolford, & Chang, 2010; J. Y. Kim et al., 2014; Wolford et al., 2013).
Histone deacetylases (HDAC) are a family of enzymes with essential roles in gene
transcriptional regulation by removal of the acetyl group of N-acetyl lysine amino acid sites on
histones. By contrast, another family of enzymes which add acetyl groups to the histone is
known as histone acetyltransferases (HAT). Hypoacetylation leads to chromatin remodeling, thus
preventing transcription factors (TF) from accessing the targeted DNA region for activation of
gene transcription. On the other hand, hyper acetylation leads to the opening of chromatin, thus
allowing more TF recruitment for increased target gene expression. Since HDACs lack intrinsic
DNA binding activity, their binding specificity and regulation of target DNA regions is largely
dependent on the direct association with transcription factors (TF), including activators or
repressors, as well as the availability of these TFs in a particular cell type (Haberland,
Montgomery, & Olson, 2009).
Because of the gene regulation function of HDACs, clinical trials with HDAC inhibitors
(HDI) are ongoing for treatment of neurodegenerative disease. A broad HDI, Vorinostat
(suberanilohydroxamic acid, abbreviated SAHA, marketed as Zolinza by Merck) was approved
17

in 2006 for treatment of persistent cutaneous T cell lymphoma (CTCL). Another HDI, Istodax
(marketed as romidepsin by Celgene) was approved in 2009 to treat CTCL and peripheral T cell
lymphoma (PTCL). To date, many other novel HDAC-based drugs are still in clinical trials or
under early stage development (Falkenberg & Johnstone, 2014).
HDACs are classified into four classes based on their sequence homologies and
evolutionary relationships. These are known as Class I (HDAC 1, 2, 3 and 8), Class IIA (HDAC
4, 5, 7 and 9), Class IIB (HDAC6 and 10) and Class IV (HDAC11) (Ruijter, Gennip, Caron,
Kemp, & Kuilenburg, 2003). Class I HDACs are ubiquitously distributed in tissues; however, the
other HDACs are mostly tissue and cell-type specific. For example, HDAC6 is found in heart,
liver, kidney, and placenta and can catalyze deacetylation of other proteins such as α-tubulin and
HSP90 in addition to histones. HDAC7 is found in heart, skeletal muscle, pancreas and placenta
and its binding partners include HIF-1α, BCL6 and endothelin receptor (Ruijter et al., 2003).
ATF3 was shown to be responsible for recruitment of several HDACs with transcription
factors to form complexes to regulate specific target gene expression. It was shown JDP2 and
ATF3 are associated with HDACs 1, 2–6 and 10 (Darlyuk-Saadon et al., 2012). Moreover,
HDAC3 and HDAC6 directly binding to JDP2 and ATF3 via their DAC conserved domains
(Darlyuk-Saadon et al., 2012). Knock- down of ATF3 and JDP2 reduce both the TSA induction
effect of histone 4 acetylation in MEF cells (Darlyuk-Saadon et al., 2012). In the macrophage,
ATF3 was reported to behave as a negative regulator of Toll-like receptor 4 (TLR4) via
interacting with HDAC1 (Mark Gilchrist et al., 2006). Interestingly, two different groups (Liu et
al., 2014; St Germain, O'Brien, & Dimitroulakos, 2010) reported that ATF3 can have synergistic
effects with HDI such as M344, cisplatin and SAHA to augment the cytotoxicity of these HDIs,
these may provide new approaches to enhance the efficiency of HDI-based tumor therapy.
18

CHAPTER 2: Hypothesis and Specific Aims
2.1. Hypothesis
We hypothesize PlGF induces HIF-1 α and ET-1 expression in endothelial cells via the
stabilization of HIF-1 α and ET-1 mRNA by post-transcriptional regulation mechanisms,
especially via miRNA. We further hypothesized that the endogenous level of miRNA candidates:
miR-648 and miR-199a are co-transcribed with their host gene. Thus, by regulating the
expression level of their host genes via different approaches (e.g., transcription factor
regulation), we expect to manipulate the endogenous level of particular miRNAs (miR-648 and
miR-199a), which may concomitantly affect their target gene activities (ET-1 and HIF-1α).

2.2. Specific Aims
Aim1: Determine whether HIF-1α and ET-1 mRNAs are post-transcriptionally regulated by
miRNAs.
Aim2: Determine the involvement of miR-199a and miR-648 in the post-transcriptional
regulation of HIF-1α and ET-1.

Aim3: Determine the transcriptional mechanisms governing the expression of miR-199a and
miR-648. Examine the potential transcription factors involved in the transcriptional regulation of
miR-199a and its host gene DNM3os, miR-648 and its host gene MICAL3.

Aim4: Determine the role of miR-199a and miR-648 in the regulation of HIF-1α and ET-1 in
mice models and correlate these results in plasma of sickle cell patients.

19

CHAPTER 3: Post-Transcriptional Regulation of Endothelin-1 by Placenta
Growth Factor via miR-648 and its transcriptional co-regulation with
MICAL3 by PAX5

3.1 Introduction
Plasma levels of ET-1 are elevated in SCA patients with PHT (Rybicki & Benjamin,
1998; Sundaram et al., 2010). The effects of ET-1 on vasoconstriction and regulation of blood
pressure have been shown utilizing ET-1 knockout mice and ET-B receptor knockout mice
(Kisanuki et al., 2010). Furthermore, ET-1 receptor (ET-R) antagonists used for treatment of
primary PHT (Benza et al., 2008) are found to be beneficial to sickle-Antilles-hemoglobin D
mice in ameliorating symptoms of PHT (Sabaa et al., 2008); thus indicating a prominent role of
ET-1/ET-receptor interaction in PHT in SCA. By contrast, the molecular mechanisms of ET-1
induction that result in sickle PHT are less well understood.
Our group previously reported that the high circulating levels of placenta growth factor
(PlGF) in SCA, as compared to healthy control subjects, correlate with increased incidence of
vaso-occlusive crises (Perelman et al., 2003). We and others (Perelman et al., 2003; Tordjman et
al., 2001) showed that PlGF is produced by erythroid cells and not by other hematopoietic cell
types. We had initially hypothesized that PlGF was selectively activating cells expressing its
cognate receptor (VEGFR1); therefore, it might be a key activator of endothelial cells and
mononuclear cells for promoting inflammation and vasoconstriction in SCA (Patel et al., 2008;
Selvaraj et al., 2003). We also observed that similar to humans with SCA, humanized sickle
20

(Berkeley-SS) mice also exhibit significantly elevated levels of PlGF and ET-1, resulting in right
ventricular (RV) hypertrophy and increased RV pressures consistent with PHT (Hsu et al., 2007;
Sundaram et al., 2010). We also demonstrated the specific contribution of elevated PlGF to PHT
by increasing erythroid expression of PlGF, using a lentivirus-vector, in normal mice to levels
seen in Berkeley-SS mice (Sundaram et al., 2010). Wild type mice, over-expressing PlGF,
consequently showed increased ET-1 production and correspondingly increased right ventricular
(RV) pressures, RV hypertrophy and pulmonary fibrosis, all consistent with features of
pulmonary hypertension (Sundaram et al., 2010). These animal findings were corroborated in
123 patients with SCA, whereupon increased plasma PlGF levels were associated with anemia,
augmented ET-1 and increased tricuspid regurgitate velocity; the latter an indirect measure of
peak pulmonary artery pressure (Sundaram et al., 2010). These studies in vivo showed that
higher plasma levels of PlGF were associated with anemia, higher levels of endothelin-1, and
clinical features of pulmonary hypertension in SCA.
We have previously shown PlGF upregulates expression of ET-1, PAI-1 and
lipoxygenase(s) in both human endothelial cells and monocytes by activation of HIF-1α,
independent of hypoxia (Patel et al., 2008; Patel, Gonsalves, Yang, Malik, & Kalra, 2009a; N.
Patel et al., 2010). In the present study, we examined the post-transcriptional mechanism of
placenta growth factor mediated endothelin-1 expression. Herein, we show the level of miR-648,
having a seed sequence complementary to the 3’-UTR of ET-1 mRNA, was attenuated in
response to treatment of cultured endothelial cells with PlGF. Moreover, we show that miR-648
located in a 5’-proximal intron of the MICAL3 gene, a member of the MICAL family of
flavoprotein monooxygenases (Grigoriev et al., 2011b), is likely co-transcribed with MICAL3
pre-mRNA and undergoes maturation following excision of the intron containing premiR-648.
21

Additionally, our studies show for the first time, to the best of our knowledge, that PAX5
transcriptionally activates co-expression of MICAL3 and pre-miR-648 in human endothelial
cells, and that the 3’UTR of ET-1 mRNA is indeed a target of miR-648. Since human miR-648
does not have a corresponding ortholog in mouse, this precluded study in animal models, e.g.
Berkeley sickle mice or PlGF-/- mice. For this reason a determination of miR-648 plasma levels
in human SCA patients was undertaken in order to corroborate the in vitro findings.

22

3.2 Materials and Methods
Reagents
Human recombinant PlGF was purchased from R&D Systems (Minneapolis, MN);
primary antibodies to endothelin-1, PAX 5 and secondary antibodies conjugated to HRP were
obtained from Santa Cruz Biotechnology (Santa Cruz, CA); antibodies against β-actin were
purchased from Sigma Chemical Company (St. Louis, MO). The PAX5 shRNA vector and
corresponding control scrambled shRNA were from Open Biosystems as a gift from Dr. Jae
Jung. Actinomycin D was purchased from Enzo Life Sciences (Plymouth Meeting, PA), and hsa-
miR mimics and hsa-miR-inhibitors were purchased from Shanghai Gene Pharma Co. Ltd
(Shanghai, China). BAC clones for ET-1 (EDN1) were obtained from Children’s Hospital
Oakland Research Institute, BACPAC Resources (Oakland, CA). The primers used for PCR
amplification of ET-1-3’-UTR and mutagenesis were purchased from Valuegene (San Diego,
CA). Plasmid pMI-PAX5 was a generous gift from Dr. Zhixin Zhang, University of Nebraska
Medical Center, Omaha, NE. Unless otherwise specified, all other reagents were purchased from
Sigma Chemical Company (St. Louis, MO).
Endothelial Cell Culture
Primary human pulmonary microvascular endothelial cells (HPMVEC) were obtained
from Cell Applications, Inc. (San Diego, CA), and human umbilical vein endothelial cells
(HUVEC) were from ATCC or Clonetics; cells were grown as per vendor's protocols. These
primary cells maintained characteristics of endothelial cell morphology and cell phenotype up to
passage 7-8, and thus were not used beyond the 8
th
passage (Patel et al., 2008). HMEC-1 cell line
was obtained from the CDC (Atlanta, GA) and cultured as described previously (K. S. Kim,
Rajagopal, Gonsalves, Johnson, & Kalra, 2006). Briefly, HMEC-1 were cultured in RPMI-1640
23

containing 10% FBS, 1 mM sodium pyruvate, 1 mM glutamine, 5 mM Hepes, MEM vitamins
and non-essential amino acids (1x), 50 g/ml endothelial cell mitogen (Biomedical
Technologies, Stoughton, MA) and heparin (20 units/ml). HMEC-1 were incubated overnight in
RPMI-1640 complete media containing 2% serum, followed by serum deprivation for 3 hrs and
subsequent treatment with either PlGF (250 ng/ml) or other experimental conditions, as
indicated.
Human Subjects
All blood samples were obtained from children with homozygous SCA at steady state
during their elective clinic appointments with routine clinical draws provided through the
Hematology Repository at Cincinnati Children’s Hospital Medical Center, Cincinnati, OH. All
samples were obtained with the informed consent of the patient/legal guardian using Institutional
Review Board approved-protocols. The plasma samples were obtained from the SCA patients
and unaffected sibling as controls (Patel, Tahara, Malik, & Kalra, 2011a).
Isolation of RNA and qRT-PCR
Endothelial cells were treated with PlGF (250 ng/ml) for indicated time periods followed
by total RNA extraction using TriZOL reagent (Invitrogen, Carlsbad, CA). Levels of mRNA and
pre-miRNA were determined and quantified using specific primers (Table 1). Real-time
quantitative PCR of mRNA and pre-miRNA templates was done using the iScript One-Step RT-
PCR Kit with SYBR Green (Bio-Rad, Hercules, CA) and an ABI Prism 7900 HT sequence
detection system (Applied Biosystems, Foster City, CA). PCR amplification of 100 ng of RNA
was performed 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, and PCR cycling and detection
24

Table 1. Oligonucleotides primers used in the Chapter 3 study.
at 95° C for 10 s, followed by 60° C for 45 s. Values are expressed as relative expression of
mRNA and pre-miRNA normalized to endogenous GAPDH mRNA (Patel et al., 2009a).

Isolation and quantification of microRNAs (miRNAs) and pre-miRNA.
Total RNA was isolated from endothelial cells using the mirVana miRNA isolation kit
(Ambion-Applied Biosystems, Foster City, CA). Specific miRNA expression was determined
25

using the TaqMan MicroRNA Assay kits for indicated miRNAs (Applied Biosystems, Foster
City, CA) as per the manufacturer’s protocol. Briefly, 100

ng of small RNA was reverse
transcribed at 16° C for 30 min, 42° C for 30 min, 85° C for 5 min and kept at 4° C. qRT-PCR (20
µ l total reactions) was performed in a 384-well plate

at 95° C for 10 min, followed by 40 cycles
of 95° C for 15 s and 60° C for 60 s. All reactions were run in triplicate. MicroRNA expression

was normalized to endogenous U6 snRNA (cell culture) or miR-16 (plasma). Pre-miRNA
expression was detected and quantified, utilizing specific pre-miRNA primers (Table I). Relative
quantitative (RQ) levels for pre-miRNA expression were calculated as 2
–ΔΔC
t

by the comparative
C
t
method, where ΔΔC
t
= (C
t
target pre-miRNA of treated sample - C
t
reference gene of treated
sample) - (C
t
target pre-miRNA of control sample - C
t
reference gene of control sample) as
previously described (Patel et al., 2011a) .
Northern Blotting
Briefly, 35 µ g of total RNA was run on a 15% non-denaturing polyacrylamide gel. The
RNA was transferred to a Biodyne B nylon membrane. The membrane was cross-linked under
UV light and pre-hybridized for 30 min using the UltraHyb hybridization buffer (Ambion, Grand
Island, NY) at 42º C. The membrane was then hybridized with biotinylated probes for miR-648
and 5S rRNA, synthesized at Valugene (San Diego, CA), at 62
o
C overnight. The membrane was
washed twice in washing buffer (Thermoscientific, Rockford, IL), followed by blocking with 5%
non-fat milk in PBS at room temperature. Streptavidin-HRP (1:250 dilutions) was added to the
membrane, incubated at room temperature for 3 hr, and followed by 2 washes with washing
buffer according to vendor’s instructions. The membranes were developed utilizing Clarity
Western ECL substrate (BioRad, Richmond, CA), and resulting images quantified using the
Image J analysis software.
26

Transient transfections
Endothelial cells (10
6
cells) were resuspended in 100 µ L of serum free RPMI 1640
medium containing indicated miRNA (60-90 pmol), anti-miRNA inhibitor (60-90 pmol),
appropriate shRNA vector and expression constructs (0.5 g), and luciferase reporter plasmids
(1.0 g), as indicated, followed by nucleofection utilizing appropriate programs in the AMAXA
nucleofector device (Lonza, Basel, Switzerland), as previously described (J. Kang et al., 2009).
The renilla luciferase plasmid (pRLSV40, 1.0 µ g) was co-transfected with firefly luciferase
reporter constructs to monitor transfection efficiency. Following nucleofection, after 24 hrs the
cells were incubated in growth medium overnight, followed by serum deprivation for 3 hrs, and
treated with PlGF (250 ng/ml) for indicated time periods. The cell lysates were assayed for
luciferase activity using the Dual Luciferase Reagent kit (Promega, Madison, WI). Luciferase
values were normalized to renilla luciferase assay values and expressed relative to the activity of
the pGL3 control vector, as appropriate. For miRNA transfections, total mRNA was extracted
using TRIzol, as described above.
Doxcycline inducible miR-648 expression plasmid
A doxycycline inducible expression plasmid was generated by PCR amplification of a
271 bp region (chr22:18463549-18463819) of MICAL3 that included the pre-miR-648 region.
The PCR fragment was inserted into the BamHI site in pLVX-EGFP vector (gift from Dr. Samad
Amini-Bavil-Olyaee), using standard cloning techniques, and designated as pLVXE-648. Correct
insertion of the sequence was determined by DNA sequencing (Retrogen, San Diego, CA).
HMEC were transfected with pLVXE-648 plasmid (1 μg/ 1x10
6
cells) by electroporation, and
transferred to complete media. After 24 hr, media was replaced with serum free media and kept
27

for 3 hr. Doxycycline (100 ng/ml) was added for indicated time periods. For washout
experiment, cells treated with doxycycline for 4 hrs, were washed 2 x in PBS, followed by PlGF
treatment for 6 hr. RNA was isolated and ET-1 mRNA was determined by qRT-PCR as
described above.
ET-1-3’-UTR plasmid and mutagenesis
The 1127 bp fragment spanning nts +974 to +2100, inclusive, relative to the translation
stop codon of ET-1 mRNA was PCR-amplified using forward and reverse primers containing
XbaI restriction enzyme sites, as listed in Table 1, and Deep Vent high fidelity DNA polymerase
(New England Biolabs, Ipswich, MA) as per standard methods. Human BAC clone for ET-1
(EDN1; NCBI NM_001168319) was used as the template (CHORI, Oakland, CA). The PCR
product was cloned downstream of the firefly luciferase gene in pGL3-Control (Promega,
Madison, WI). The orientation and identity of the insert relative to the luciferase gene was
confirmed by DNA sequencing and the plasmid was purified using EndoFree Plasmid Maxi Kit
(Qiagen, Valencia, CA). The resulting plasmid is designated pGL3-ET-1-3’-UTR. Mutation of
the miR-648 binding sites in the ET-1 3’UTR was generated using pGL3-ET-1-3’-UTR as the
template with primers listed in Table 1 in accordance with the NEB Q5 site-directed mutagenesis
protocol (New England Biolabs, Ipswich, MA). The mutations were confirmed by DNA
sequencing (Retrogen, La Jolla, CA).
Western Blots
Endothelial cells (10
6
cells) were incubated in serum-free media overnight, followed by
treatment with PlGF for the indicated time periods. Protein lysates were prepared as previously
described and subjected to electrophoresis. Samples were transferred to a PVDF membrane
(BioRad, Hercules, CA). Membranes were probed with antibodies (diluted as indicated) for
28

PAX5 (1:250) and ET-1 (1:500). Membranes were stripped and re-probed with an antibody to β-
actin (1:50,000), to normalize any loading differences. Protein bands were detected using the
West Pico Chemiluminescent substrate (Thermo Scientific) and sizes were compared to pre-
stained molecular weight markers (NEB, MA).
Quantitation of transcripts by semiquantitative RT-PCR
Total RNA was extracted from HMEC-1 using TriZOL reagent as previously described.
RNA transcripts were reverse-transcribed and amplified using specific primers for P1, P2, P2a
products and GAPDH (Table 1) utilizing the Access RT-PCR system (Qiagen, La Jolla, CA).
Briefly, 200 ng (non-limiting) or 25 ng (limiting) total RNA was reverse transcribed at 45° C for
45 min to generate cDNA products followed by heat-inactivation at 94° C for 2 min. PCR
amplification was performed as follows: 94° C for 30 s, 60° C for 1 min and 68° C for 2 min for 40
cycles, and a final extension at 68° C for 7 min. The PCR products were run on a 2% agarose gel,
visualized by ethidium bromide and quantitated by Image J software analysis.
Chromatin Immunoprecipitation assays (ChIP)
HMEC cells (10
7
cells) were serum starved prior to treatment with PlGF as described
above. Chromatin immunoprecipitation was performed using PAX5 antibody as previously
described (Patel et al., 2008). Immuno-precipitated DNA was subjected to PCR for 30 cycles, as
follows: 94° C for 1 min, 60° C for 1 min and 72° C for 1 min, using the primers specified in Table
I. PCR products were analyzed by agarose gel electrophoresis and compared to DNA size
markers.
Statistical analysis
Data are presented as means ± SEM. Student t-test was used to evaluate the significance
of differences between paired samples. SCA vs. control plasma samples for qRT-PCR were
29

analyzed utilizing unpaired t-test. Values of P < 0.05 were considered significant. Statistical
significance as indicated in the figures: ***P<0.001, **P<0.01, *P<0.05 and ns, not significant.

30

3.3 Results
Post-transcriptional regulation of PlGF-mediated ET-1 mRNA expression
Previous studies from our lab show that in the endothelial cells, PlGF induces ET-1
mRNA in a time-dependent manner, reaching a maximum level at 6 hours post-induction, with
declining levels persisting through 24 hours (Patel et al., 2008).
To demonstrate the possible post-transcriptional regulatory mechanism of ET-1 in
HMEC-1, we utilized actinomycin D. As shown in Figure 4A, PlGF significantly stabilized ET-1
mRNA, increase of t
½

of ET-1 mRNA from 22 ± 1 min to 36 ± 2 min. The decay kinetics of ET-
1 mRNA in HMEC-1 with actinomycin D treatment (10μg/mL) in the absence of PlGF showed a
biphasic decline, consistent with a second order degradation process, possibly a miRNA-
dependent mechanism.
31

Figure 4. PlGF-mediated expression of candidate miRNAs in human endothelial cell line (HMEC-1) with
predicted binding site(s) within the 3’UTR of ET-1 mRNA. A. Determination of ET-1 mRNA half-life (t
1/2
)
in response to PlGF treatment. B. Schematic showing 3’UTR of ET-1 mRNA with binding site(s) for
selected miRNAs. C. HMEC-1 was treated with PlGF for 6 hrs, followed by isolation of miRNAs and
quantitation by qRT-PCR. The relative expression levels of indicated miRNAs were normalized to U6
snRNA. D. HMEC-1 were treated with PlGF (250 ng/ml) for 6 hrs followed by mRNA isolation and
quantitation of pre-miRs by qRT-PCR. Results are expressed as fold change between PlGF-treated vs.
untreated sample.

Identification of miRNAs involved in destabilization of ET-1 mRNA in HMEC-1
To identify the potential miRNA candidates involved in the ET-1 mRNA destabilization,
bioinformatic analysis (URL: http://www.ebi.ac.uk/enright-srv/microcosm/cgi-
bin/targets/v5/detail_view.pl?transcript_id=ENST00000379375) was performed to identify
miRNA candidates which target ET-1 mRNA. The analysis is based on the degree of ET-1
32

mRNA complementarity to specific miRNAs, as quantified by the Gibbs Energy of seed
sequence stability ( G° ) and evolutionary conservation among mammalian species of miRNA
target sites within the ET-1 3’-UTR. Based these two criteria, we selected three candidates for
further study, and showed them in the schematic shown in Fig.4B (miR-648, miR-517B, and
miR-934). In addition, we also included miR-125a-3p, miR-125a-5p, and miR-125b, since they
are reported to be involved in oxidized LDL-mediated regulation of ET-1 mRNA in vascular
endothelial cells (D. Li et al., 2010). Lastly, we also included miR-199a since the stability of
HIF-1α mRNA is affected by miR-199a in response to ethanol (Yeligar, Machida, & Kalra,
2010).
The levels of the selected miRNAs were determined in HMEC-1 cells in response to
PlGF treatments. As shown in Fig.4C, after PlGF treatment, miR-934 and miR-648 levels were
decreased by ~3-fold and ~16-fold, respectively; similarly, miR-199a-5p level decreased by ~8-
fold. However, the level of miR-199a-3p did not change significantly in response to PlGF
(Fig.4C). miR-125a-3p, miR-125a-5p, and miR-125b increased modestly in the range of 2- to 4-
fold as shown in the Fig.4C.
Since mature miRNAs are derived from pre-miRNAs and cleaved by Dicer in the cytoplasm
(REF), we examined whether a change of pre-miR-648 and pre-miR-934 level in the presence of
PlGF were correlated with the observed change in respective mature miRNAs. As shown in Fig.
4D, PlGF decreases pre-miR-648 and pre-miR-934 levels by ~80% and ~75%, as detected by the
qTR-PCR, respectively, which are consistent with the attenuated level of mature miR-648 and
miR-934 level in HMEC-1 in response to PlGF. In summary, these data showed that PlGF was
responsible for decreased transcription of miR-648 and miR-934 in HMEC-1.

33

Effect of miRs and anti-miRs on ET-1 mRNA level in human endothelial cell line, HMEC-1
Since miR-648 levels decreased ~16-fold in PlGF treated HMEC-1 (Fig. 4C), which gave
the most significant experimental result among the candidate miRNAs, we focused our studies
on this particular miR-648. First, we examined whether miR-648 directly affected the level of
ET-1 mRNA under basal and PlGF-treated conditions by utilizing miR-648 mimic, the artificial
oligonucleotide of mature miR-648. As shown in Fig. 5A, transfection of HMEC-1 with two
concentration of miR-648 mimic, 60 pmol and 90 pmol, dose-dependently attenuated the basal
level of ET-1 mRNA by ~40% and ~75%, respectively (Fig. 5A, lane 2 vs. lane 1). Alternatively,
transfection of 60 pmol and 90 pmol of anti-miR-648, the artificial single strand
complementarily oligonucleotide of miR-648, antagonized endogenous miR-648 and augmented
the levels of ET-1 mRNA by ~2-fold and ~2.5-fold, respectively (Fig. 5A, lane 3 vs. lane 1). To
validate the above results, we utilized negative controls, scrambled (sc) miR siRNA (sc mimic)
and anti-miR-siRNA (sc inhibitor). Both of these did not affect ET-1 mRNA expression (Fig.
5A, lanes 4 and 5 vs. lane 1).
Moreover, we transfected miR-648 mimic at both 60 pmol and 90 pmol into HMEC-1.
Data showed that the transfection of miR-648 mimics reduced ET-1 mRNA expression by ~65%
and ~95%, respectively, relative to PlGF-treatment alone (Fig. 5A, lane 7 vs. lane 6).
Conversely, transfection with 60 pmol and 90 pmol of anti-miR-648, followed by PlGF
treatment, antagonized endogenous miR-648, and resulted in increased levels of ET-1 mRNA by
~38% and ~45%, respectively, over and above PlGF treatment alone (Fig. 5A, lane 8 vs. lane 6).
34

Figure 5. Effect of exogenous miR-648 on ET-1 in HMEC-1. A. ET-1 mRNA expression in cells
transfected with miRs and anti-miR under basal and PlGF-treated condition. B. Western blot of HMEC-1
cell transfected with either miR-648 mimic (90 pmole) or anti-miR (90 pmole). C. Western blot of HMEC-
1 cell lysates transfected with either miR-648 mimic or anti-miR following PlGF treatment for 24 hr. D
and E. Northern blot for miR-648 and 5S rRNA. F. ET-1 mRNA expression in cells transfected with
pLVX-EGFP and pLVXE-648 plasmids.

35

Previous results showed that miR-648 attenuates ET-1mRNA, however the effect of miR-
648 on ET-1 translation is not known. To address this, ET-1 protein level were examined by
western blotting in HMEC-1 transfected with miR-648 mimic and antimiR-648. There was
~50% reduction of ET-1 protein (Fig. 5B, lane 2 vs. lane 1) after transfection with miR-648
mimic, and anti-miR-648 increased expression of ET-1 protein around 2-fold (Fig. 5B, lane 3 vs.
lane 1) under basal conditions (n=3). The effect of exogenous miR-648 treatment on ET-1
protein level was also examined in the presence of PlGF. PlGF induced ET-1 protein levels by 2-
fold (Fig. 5C, lane 2 vs. lane 1); however, miR-648 with PlGF treatment, abrogated induction of
ET-1 protein by ~70% (Fig. 5C, lane 3 vs. lane 2) and anti-miR-648 had no effect on ET-1
protein level (Fig. 5C, lane 4 vs. lane 2). These results indicate that exogenous miR-648 reduces
basal levels and induced level of ET-1 mRNA, and concomitantly reduces ET-1 protein levels.
Next, we performed Northern blots utilizing biotinylated miR-648 probe to detect the
expression level of endogenous miR-648 in HMEC-1. As shown in Fig. 5D, transfection of miR-
648 mimic in HMEC-1 increased ~3-fold endogenous levels of miR-648, while anti-miR-648
completely antagonized endogenous levels of miR-648 (lane 3 vs. lane 1).
Since endogenous levels of miR-648 may vary between endothelial cells derived from
primary culture vs. established cell lines, we also quantified the expression of miR-648 in these
cells by Northern blotting. As shown in Fig. 5E, both HUVEC and HMEC-1 showed almost
equivalent amounts of miR-648, while HPMVEC showed ~2.6 fold higher expression of miR-
648. Thus, these results showed miR-648 is constitutively expressed in HMEC-1 and it suggests
that miR-648 is responsible for maintenance of low ET-1 levels by post-transcriptional
repression.
36

Next, a doxycycline (dox) inducible pre-miR-648 plasmid was transfected into HMEC-1
to validate the effect of miR-648 on ET-1 mRNA. As shown in Fig. 5F, dox-induction of
HMEC-1 showed that time dependent (0.25-4 hr) miR-648 synthesis, which directly correlated
with a decline in ET-1 mRNA expression levels in HMEC-1 cells (Fig. 5F, lanes 5, 6 and 7 vs.
lane 4). Dox-induction of control vector transfected cells showed no effect in ET-1 mRNA level
(Fig. 5F, lane 3 vs. lane 1). Moreover, after dox was washed out by PBS, miR-648 induction is
reversed by PlGF treatment, which restored to basal ET-1 mRNA expression (Fig. 5F, lane 8 vs.
lane 4). Taken together, these data confirmed that miR-648, expressed in situ, repressed the
expression of ET-1 mRNA and protein in HMEC-1. Thus physiological reduction of miR-648
levels in response to PlGF should positively contribute to expression of ET-1.

miR-648 regulates ET-1 mRNA and protein expression in primary human endothelial cells
We examined whether miR-648 played a role in the regulation of ET-1 in primary human
endothelial cells. Treatment of human umbilical vein endothelial cells (HUVEC) and human
pulmonary microvascular endothelial cells (HPMVEC) with PlGF resulted in ~70% reduction in
miR-648 expression (Fig. 6A).
Next, the relationship between miR-648 levels to ET-1 induction was examined in these
primary endothelial cells following treatment with PlGF. Transfection of HUVEC with miR-648
mimic reduced endogenous ET-1 mRNA levels by ~50% (Fig. 6B, lane 2 vs. lane 1).
Conversely, anti-miR-648 augmented ET-1 mRNA by ~2.2 fold compared to untreated cells
(Fig. 6B, lane 3 vs. lane 1). Next, we examined whether miR-648 modulated ET-1 expression
following PlGF treatment. Incubation of HUVEC with PlGF for 6 hrs resulted in ~2.5-fold
37

induction of ET-1 mRNA level (Fig. 6B, lane 6 vs. lane 1), as previously shown (Patel et al.,
2008). Transfection of HUVEC with miR-648 mimic, followed by PlGF treatment reduced ET-1
mRNA levels to the basal level (Fig. 6B, lane 7 vs. lane 6). Conversely, transfection of anti-miR-
648 in the same condition did not change ET-1 mRNA expression beyond PlGF-treatment alone
(Fig. 6B, lane 8 vs. lane 6). Western blotting showed that in the presence of PlGF, transfection of
HUVEC with miR-648 mimic attenuated ET-1 protein to the basal level (Fig. 6C, lane 3 vs. lane
2). However, anti-miR-648 did not significantly increase ET-1 protein levels over and above that
observed with PlGF plus sc-anti-miR (Fig. 6C, lane 4 vs. lane 6) and also not above PlGF-
treatment alone (Fig. 6C, lane 4 vs. lane 2). Moreover, in PlGF-treated human pulmonary
endothelial cells (HPMVEC), miR-648 mimic effectively reduced ET-1 mRNA expression (Fig.
6D, lane 3 vs. lane 2) while anti-miR-648 did not significantly affect ET-1 mRNA levels over
and above PlGF-treatment alone (Fig. 6D, lane 4 vs. lane 2). In summary, these data showed that
miR-648 was regulating ET-1 mRNA and protein levels both under basal and PlGF treated
conditions in primary human endothelial cells, same as HMEC-1, thus contributing to permissive
conditions for ET-1 expression.
38

Figure 6. Endogenous miR-648 expression and effects of exogenous miR-648 and anti-miR-648 on ET-
1 mRNA and protein in cultured primary human endothelial cells. A. miR-648 expression in PlGF-treated
HUVEC and HPMVEC. B. ET-1 mRNA expression in HUVEC transfected with miR and anti-miR under
basal and PlGF treated (6hrs) condition. C. Western blot for expression of ET-1 protein in HUVEC
transfected with indicated miR or anti-miR, followed by PlGF treatment for 12 hr. D. Effect of miR-648
and anti-miR-648 on ET-1 mRNA in HPMVEC.

39

miR-648 directly interacts with predicted target site in the 3’-UTR of ET-1 mRNA
To address the question whether miR-648 regulated ET-1 via direct interacting with the
3’-UTR, we generated a luciferase reporter construct in which the 3’-UTR of ET-1 mRNA
(bases +2 to +1174, relative to the stop codon) region, which contains the putative miR-648
binding site, was inserted downstream of the luciferase open reading frame (Fig. 7A). This
construct was named as pGL3-ET-1-3’UTR, and it has the predicted miR-648 binding site.
HMEC-1 cells were then transfected with this luciferase reporter. Upon PlGF treatment, there is
a ~3-fold increase in luciferase reporter activity, indicating that this luciferase mRNA can
respond to PlGF like native ET-1 mRNA (Fig. 7B, lane 5 vs. lane 2). Co-transfection of pGL-3–
ET-1-3’UTR reporter with miR-648 mimic (90 pmol) modestly reduced luciferase activity under
basal conditions (Fig. 7B, lane 3 vs. lane 2). However, miR-648 decreased the luciferase activity
by ~25% in cells treated with PlGF (Fig. 7B, lane 6 vs. lane 5). The specificity of miR-648 on
reporter mRNA activity was elucidated by utilizing anti-miR-648. The effects of endogenous
miR-648 (previously demonstrated in Figs. 5A and 2B) were antagonized by anti-miR-648 under
both basal (Fig. 7B, lane 4 vs. lane 2) and PlGF treatments (Fig. 7B, lane 7 vs. lane 5). In both
conditions, anti-miR-648 increased the luciferase reporter activity by ~2-fold.
40

To consolidate our previous results, we performed site-directed mutagenesis in the
corresponding miR-648 seed sequence of the pGL3-ET-1-3’-UTR reporter (Fig. 7A), which is
referred to as pGL3-ET-1(mut)-3’-UTR. As shown in Fig. 7C, co-transfection of both miR-648
Figure 7. Effects of miR-648 and anti-miR-648 oligonucleotides on ET-1-3’UTR reporter luciferase
activity. A. Schematic representation of the ET-1-3’UTR luciferase reporter construct. The nucleotide
sequence of the predicted miR-648 binding site within the ET-1-3’-UTR is shown as wild type (wt); point
mutations in the seed sequence of the mutant (mut) are indicated by asterisks. B. HMEC-1 were co-
transfected with wt ET-1-3’-UTR plasmid and the indicated miR or anti-miR (90 pmole) followed by 6 hr
PlGF treatment, 24 hr post-transfection. The luciferase activity was normalized to renilla activity. C.
HMEC were transfected with mutant ET-1-3’UTR reporter plasmid and the indicated miR-648 mimic or
anti-miR-648.

41

mimic and anti-miR-648 with the mutant reporter did not significantly affect basal luciferase
expression (Fig. 7C, lane 3 vs. lane 2, and lane 4 vs. lane 2). Taken together, these data showed
that miR-648 directly targeted the 3’-UTR of ET-1 mRNA for turnover, and subsequently down
regulated the expression of ET-1.

Transcription of miR-648, located in an intron of MICAL3, is co-regulated with MICAL3
miR-648 gene is located in the first intron of MICAL3, a member of the MICAL family
of flavoprotein monooxygenases, (Hung et al., 2010; Kolk & Pasterkamp, 2007). The
orientation and location of the miR-648 gene suggests that it is transcribed from the same DNA
template strand as MICAL3, thus we hypothesized that pre-miR-648 is within a MICAL3
transcription unit and co-transcribed (Fig. 8A). There are at least three functional promoters (P1,
P2 and P3) associated with the MICAL3 locus (Fig. 8A) and at least 19 possible primary
transcripts (ENSEMBL). Not all of the primary transcripts encode a protein, thus suggesting
these may serve some other physiological function (J. T. Lee, 2012).
42

Our previous results revealed that miR-648 destabilizes ET-1 mRNA and PlGF signaling
Figure 8. Role of P1, P2 and internal P2a promoter in transcription of MICAL3. A. Schematic of 5’-region of
MICAL3 showing promoters P1, P2 and internal promoter P2a; location of 5’-proximal exons and intronic
miR-648. Lower lines indicate splicing patterns of transcripts initiated at either P1 (distal) or P2 (medial)
promoters. B. Effect of PlGF treatment on internal promoter P2a luciferase activity. Cells were transfected
with pGL3-P2a luc plasmid, followed by treated with PlGF for indicated time periods. C. Quantitation of P1,
P2 and P2a transcripts and GAPDH transcript in untreated and PlGF treated cells, as analyzed by gel
electrophoresis and ethidium bromide staining.
43

causes decreased miR-648 synthesis in endothelial cells, a further examination showed that
MICAL3 distal (P1) and medial (P2) promoters can be responsible to generate miR-648, thus we
examined whether PlGF treatment of endothelial cells affected these potential promoters for
miR-648 transcription.
Since there is a large distance between the distal promoter (P1) transcription start site and
miR-648, it was possible that a cryptic promoter proximal to miR-648 was actually responsible
for miR-648 synthesis. In order to answer this question, we inserted the ~2kb DNA fragment, 5’-
adjacent to miR-648, into pGL3-Basic luciferase reporter, named pGL3-P2a, to determine the
presence of a potential internal promoter. As shown in Fig. 8B, the pGL3-P2a construct
exhibited a strong, 6-fold expression of luciferase activity in HMEC-1 cells compared to the
promoter-less pGL3-Basic vector (Fig. 8B lane 2 vs. lane 1). This indicated the ~2kb DNA
fragment inserted into pGL3-P2a is a functional DNA region with promoter activity. Then, this
promoter (provisionally named P2a) was tested for any change in activity in response to PlGF. In
the presence of PlGF, HMEC-1 cells transfected with pGL3-P2a showed no change in luciferase
activity from 1-6 hr compared to the untreated control (Fig. 8B, lanes 3-6 vs. lane 2). Since a
strong reduction was observed of miR-648 level in response to PlGF (Fig. 4C), it is unlikely that
P2a is responsible for this reduction. Another possible explanation can be the repressive
element(s) were excluded from the ~2 kb DNA segment inserted.
To determine the relative transcription activities of P1, P2 and P2a, a semi-quantitative
RT-PCR experiment of endogenous RNA was performed with primers targeting the first exons
of P1, P2 and P2a RNA products. With limiting RT-cDNA substrate, P1 RNA product showed
the strongest signal (product size of 183 bp), while P2 showed modest signal (product size of 102
bp), and P2a RNA did not yield any products (product size of 173 bp) (Fig. 8C). Quantification
44

of the signal showed that P1 promoter was ~12-fold more active than P2 promoter under basal
condition. Moreover, the P1 RNA product showed a ~45% reduction after PlGF treatment
compared to the untreated control (Fig. 8C, lane 6 vs. lane 2). GAPDH is used as an internal
control (Fig. 8C, lanes 5 and 9) to ensure any changes were not the result of uneven sample
handling or PCR analysis. Taken together, we can conclude that P1 is the predominant promoter
controlling transcription of miR-648, and promoters P2 and P2a were not responsible for the
bulk of miR-648 synthesis compared to P1 in HMEC-1 cells.
45

A bioinformatics analysis (TRANSFAC) of the MICAL3 locus revealed the presence of
several transcription factor binding motifs, e.g. sites for PAX5, SMAD3, HNF3A and HNF4A,
upstream of the P1 promoter. Among these transcription factors, we selected PAX for further
analysis, since there are three putative binding sites for PAX5 (Fig. 9A) nearest the P1-TSS of
MICAL3.
Figure 9. Expression of PAX5 in endothelial cells. A. Schematic showing PAX5 binding sites in the 5’
region of MICAL3-P1 promoter. B. Western blots of cell lysates from HMEC-1, HUVEC, HPMVEC and B-
cells, showing expression of PAX5 protein. C. D. and E. Effects of silencing and gain of function of PAX5
on PAX5 mRNA and protein. HMEC-1 was transfected with PAX5 shRNA or expression plasmid pMI-
PAX5.

46

We first determined the presence of PAX5 in different endothelial cells. Western blot
analysis of cell lysates obtained from HMEC-1, HUVEC and HPMVEC were all positive for
expression of PAX5 protein. Moreover, a B cell lysate was used as a positive control for this
factor (Fig. 9B) since PAX5 was mainly reported to involve in B cell lineage differentiation
(Cobaleda, Schebesta, Delogu, & Busslinger, 2007; He et al., 2011). Besides of B cells, PAX5 is
also observed in other hematopoietic cells, cancers and neural cell types (Gerard et al., 1995;
O'Brien, Morin, Ouellette, & Robichaud, 2011; G. J. Zhao et al., 2012).
To examine whether PAX5 is involved in MICAL3 transcription regulation, we
performed loss and gain of PAX5 function in HMEC-1. Transfection with PAX5 shRNA
effectively reduced endogenous PAX5 mRNA levels (Fig. 9C, lane 2 vs. lane 1), while
transfection of the exogenous gene copy PAX5 (pMI-PAX5) into HMEC-1 increased
intracellular levels of PAX5 mRNA (Fig. 9C, lane 4 vs. lane 1). Moreover, PAX5 shRNA
reduced PAX5 protein levels by ~50% (Fig. 9D, lane 3 vs. lane 1); while PAX5 expression
plasmid augmented PAX5 protein levels by ~170% compared to untreated cells (Fig. 9E, lane 3
vs. lane 1). These results clearly showed PAX5 is present in endothelial cell line (HMEC-1)
under basal conditions.

miR-648 is co-transcribed with MICAL3-P1 transcript, and regulated by PAX5
transcription factor
As discussed above, transcription from P1 was responsible for miR-648 synthesis. Thus,
the next question is whether this P1 promoter is regulated by PAX5. In the absence of PlGF, P1
transcript(s) were significantly reduced (~60%) by PAX5 shRNA (Fig. 10A, lane 2 vs. lane 1),
47

while overexpression of PAX5, utilizing pMI-PAX5 plasmid, increased P1 utilization by ~3.5
fold relative to control (Fig. 10A, lane 4 vs. lane 1). Transfection with either control scRNA or
pMI empty vector did not significantly affect levels of MICAL3 P1 promoter transcript(s) (Fig.
10A, lanes 3 and 5 vs. lane 1).
Furthermore, we examined whether PAX5 transcriptionally regulated expression of pre-
miR-648 in cultured human endothelial cells. We assumed that splicing of MICAL3 pre-mRNAs
was an obligatory event leading to pri-miR-648 and pre-miR-648 synthesis, barring any
regulation of post-transcriptional processing or nuclear export of the pre-miRNA. Transfection of
HMEC-1 with PAX5 shRNA reduced pre-miR-648 expression by >90% (Fig. 10A, lane 2 vs.
lane 1) while PAX5 overexpression vector (pMI-PAX5) augmented pre-miR-648 by >2.5 fold
(Fig. 10A, lane 4 vs. lane 1). Transfection of HMEC-1 with either sc-RNA or empty vector
(pMI) had no significant effect on pre-miR-648 expression (Fig. 10A, lanes 3 and 5 vs. lane 1).
These data showed that PAX5 was required for initiating transcription from the distal (P1)
promoter of MICAL3, which is the key process of generating pre-miR-648.
48

Next, we examined whether the same conclusion is also true in primary human
Figure 10. PAX5 regulates transcription from MICAL3-P1 promoter and expression of pre-miR-648. A.
and B. qRT-PCR of HMEC-1(A) or HUVEC(B) transfected with PAX5 shRNA and pMI-PAX5 expression
plasmid, or scramble RNA. C. PlGF treatment of HMEC reduces PAX5 protein level. D. Treatment of
HMEC with PlGF reduced expression of PAX5 and MICAL3-P1 promoter transcripts, which was further
reduced by PAX5 shRNA. E. and F. In vivo association of PAX5 with PAX5 sites proximal to P1

49

endothelial cells. Similar to HMEC-1, transfection of HUVEC with PAX5 shRNA attenuated
pre-miR-648 synthesis and MICAL3-P1 promoter transcripts (Fig. 10B) and pMI-PAX5
augmented pre-miR-648 synthesis and MICAL3-P1 promoter activity (Fig. 10B). These data
clearly showed that PAX5 was present in primary human endothelial cells (HUVEC) and was
functional in regulating expression of both MICAL3 and pre-miR-648 in primary human
endothelial cells.

PlGF attenuates PAX5 levels and associated transcription from MICAL3-P1 promoter
PlGF decreased PAX5 protein expression ~70% as shown by western blot of HMEC-1
lysates (Fig. 10C), which correlated with a ~65% decrease PAX5 mRNA expression (Fig. 10D,
lane 2 vs. lane1). The decreased expression of PAX5, in the presence of PlGF, correlated with
~70% reduction of transcription activity from MICAL3-P1 (Fig. 10D, lane 2 vs. lane 1); as
shown before, under these conditions, transcription of pre-miR-648 was reduced ~80% (Fig.
4D). Furthermore, PAX5 shRNA followed by PlGF treatment further reduced PAX5 mRNA
levels (Fig. 10D, lane 3 vs. lane 2), while scRNA (non-specific shRNA) had no such effect (Fig.
10D. lane 4 vs. lane 2). Thus we can conclude that PAX5 levels were reduced significantly by
PlGF, and concomitantly reduced levels of MICAL3-P1 transcript(s) and pre-miR-648, thus
indicating PAX5 is an important positive regulator of MICAL3 and miR-648 transcription, and
needed for basal level expression.

50

PAX5 binds to native chromatin for transcription of MICAL3 as demonstrated by
chromatin immunoprecipitation (ChIP) analysis
To understand the mechanism of PAX5 regulation of MICAL3 transcription, we utilized
chromatin immunoprecipitation (ChIP) analysis of the potential PAX5 binding locus on
MICAL3 promoter in uninduced HMEC-1. These cells were also transfected with either PAX5
shRNA or pMI-PAX5 and subjected to the same ChIP analysis. The predicted PAX5 binding
sites are located at bases -1327 to -1323 (site #1), bases – 2247 to -2243 (site #2), and bases -
2312 to -2308 (site #3). Site #1 can be detected independently (as amplicon #1), whereas sites #2
and #3 are too close together to be resolved by PCR amplification (Fig. 9A). For this reason,
amplicon #2 (bases -2331 to -2238) will detect occupancy of either site #2 or #3 or both. As
shown in Figs. 10E and 10F, this analysis showed that PAX5 was present on the MICAL3
promoter under basal conditions, based on recovery of amplicon #1 (100 bp product) and
amplicon #2 (94 bp product) (Fig. 10E, lane 1 and Fig. 10F, lane 1, respectively). Knockdown of
PAX5 by PAX5 shRNA completely abrogated recovery of amplicon #1 and #2 (Fig. 10E, lane 2
and Fig. 10F, lane 2, respectively), whereas cells overexpressing PAX5 (pMI-PAX5) showed
~70% and ~125% increases in amplicon #1 and #2 PCR products (Fig. 10E, lane 3 and Fig. 10F,
lane 3, respectively). Chromatin immunoprecipitated with non-specific IgG did not yield a
MICAL3-specific PCR product (Figs. 10E and 10F, bottom panels). Furthermore the input DNA
was equivalent in control, PAX5 shRNA and pMI-PAX5 transfected chromatin samples (see
middle panels of Figs. 10E and 10F). Together, these data showed that PAX5 binding, proximal
to P1, is correlated with basal MICAL3 and pre-miR-648 transcription in endothelial cells.

51

miR-648 levels in plasma from SCA patients
Our previous studies showed that SCA patients have a higher level of PlGF in plasma
compared to the healthy subjects, and significantly correlated with increased plasma ET-1 levels
and tricuspid regurgitant velocity; the latter is reflective of peak pulmonary artery pressure or an
indicator of PHT (Sundaram et al., 2010). It is pertinent to mention that human miR-648 does not
have a corresponding ortholog in mouse, this precluded studies in animal models, e.g. Berkeley
sickle mice or PlGF
-/-
mice. Since miR-648, in the present study, post-transcriptionally
modulated ET-1 expression, we measured plasma levels of miR-648 in SCA patients along with
their unaffected sibling controls. The results showed circulating levels of miR-648 were
detectable in samples from both SCA patients (n=13) and controls (n=13). However, miR-648
plasma levels were statistically different between SCA patients compared to unaffected controls
(mean ± SEM of 0.39 ± 0.10 in SCA vs. 1.54 ± 0. 50 in AA; p= 0.034) as indicated in Fig. 11A.
The significantly lower plasma levels of miR-648 in SCA patients are in agreement with the
relationship between miR-648 and ET-1 observed in vitro.
Figure 11. miR-648 plasma levels in SCA and control subjects, and working model of miR-648 mediated
regulation of ET-1 expression. A. The plasma concentrations of miR-648 were quantified from SCA
patients (n=13) and healthy matched controls (n=13). The relative expression levels of miRNAs in these
samples were normalized to endogenous miR-16. B. Working model of miR-648 mediated regulation of
ET-1 expression via its binding to the 3’UTR of ET-1 mRNA.

52

3.4 Discussion
This study extends our understanding of the regulatory mechanisms underlying
expression of endothelin-1 and its association with pulmonary hypertension in sickle cell disease.
PlGF treatment of endothelial cells resulted in a net stabilization of ET-1 mRNA from the basal
state and the mechanism for this change exhibited characteristics of a miRNA based process. For
this reason we examined whether expression of specific miRNAs predicted to bind to the 3’-
UTR of ET-1 mRNA changed in endothelial cells upon PlGF treatment. From seven candidate
miRNAs as identified by bioinformatics analysis as potential effectors of ET-1 mRNA stability,
we found that PlGF treatment significantly reduced the expression of miR-648. And we
hypothesized that changes in endogenous miR-648 levels would be expected to have a
significant effect on cytoplasmic ET-1 mRNA levels.
We then tested the post-transcriptional regulation of ET-1 by miR-648. The specific
decreased miR-648 in response to PlGF was linked to increased stabilization of ET-1 mRNA.
This link was confirmed and extended using miR-648 mimic to decrease endogenous ET-1 under
basal conditions and during PlGF induction. The anti-miR-648 abrogated basal turnover of ET-1
mRNA, demonstrating the importance of miR-648 in suppressing activity of this potent
vasoconstrictor under these conditions. Finally, we demonstrated using a luciferase translation
reporter that the primary sequence required for regulating ET-1 mRNA stability did indeed
reside in the 3’-UTR of this mRNA. The synthetic luciferase reporter fused to the wt. ET-1
3’UTR behaved exactly like native ET-1 mRNA under basal and PlGF induction in endothelial
cells. By contrast, mutation of the miR-648 recognition element in the 3’-UTR of the reporter
completely nullified any effect of endogenous miR-648 or the synthetic mimic.
53

The synthesis of miR-648 afforded new insights into how this regulatory RNA is itself
regulated at the level of transcription. The miR-648 gene is present in the first intron of
MICAL3, a member of the microtubule associated monooxygenase, calponin and LIM domain
containing (MICAL) family of flavoprotein monooxygenases, which participate in axon
guidance, actin remodeling (Hung et al., 2010; Terman, Mao, Pasterkamp, Yu, & Kolodkin,
2002), and redox activity in promoting vesicle-docking complexes in the process of exocytosis
(Grigoriev et al., 2011a). We asked whether the precursor of this miRNA was derived from
MICAL3 transcripts or arose from an independent transcription unit driven by an independent
promoter. Although the MICAL3 locus is regulated by at least three promoters, giving rise to 19
total, alternatively spliced transcripts (ENSEMBL), our results showed that the P1 (distal)
promoter of MICAL3 was regulated and responsible for expressing the RNA precursor of miR-
648. Although we determined that the 5’-flanking, 2 kb intron region (P2a) adjacent to miR-648
showed promoter activity with a heterologous luciferase reporter, it was not regulated by PlGF.
Moreover in the context of the MICAL3 locus, this potential promoter did not show appreciable
activity. Thus, we are confident that the P1 promoter is responsible for miR-648 synthesis.
We identified PAX5 as a transcription factor involved in the co-transcriptional regulation
of MICAL3 and miR-648 utilizing loss and gain of PAX5 function approaches in endothelial cell
line HMEC-1. The requirement for PAX5 in basal MICAL3 transcription was demonstrated by
transfection of HMEC-1 and HUVEC with PAX5 shRNA, wherein the latter attenuated
MICAL3 mRNA and miR-648 expression. Conversely, constitutive expression of PAX5 with
exogenous PAX5 augmented MICAL3 transcription indicating that PAX5 was indeed functional
in regulating transcription of this gene in endothelial cells.
54

These above studies were extended by demonstrating enrichment of PAX5 binding to the
proximal 2.5 kb segment upstream of the MICAL3 P1 promoter, as shown by ChIP analysis. We
actually identified seven potential PAX5 sites based on the consensus PAX5 binding motif,
upstream of MICAL3 P1 promoter. However as the present study demonstrated, the three PAX5
sites nearest the TSS were sufficient to retain regulation by PlGF. Further studies will be needed
to establish how PAX5 activity is down regulated in the context of PlGF induction and
identification of additional bona fide PAX5 binding sites proximal to P1.
It was also of interest to learn that PAX5 protein is expressed in human primary
endothelial cells, as detected by western blot analysis of transformed human endothelial cell line
(HMEC-1), human umbilical vein endothelial cell line (HUVEC) and human pulmonary
microvascular endothelial cells (HPMVEC). Although PAX5 is well-known as a B cell lineage
specific activator protein and required in B cell lineage development (Cobaleda et al., 2007; He
et al., 2011), it is in fact expressed in almost all cell types (Su et al., 2004). Thus it would be of
great interest to determine how PAX5 expression is regulated in endothelial cells in response to
PlGF.
Our studies on ET-1 offer a second example of a bipartite mechanism for gene induction,
analogous to one we showed for PAI-1 (39). For the latter we observed an immediate down
regulation of miRNAs responsible for suppressing basal PAI-1 mRNA levels, which results in an
early burst of PAI-1 synthesis. A sustained increase in PAI-1 expression was achieved by a
slower HIF-1α induction of the PAI-1 gene with corresponding appearance of newly transcribed
PAI-1 mRNA in the cytoplasm. In the ET-1 system, down-regulation of miR-648 apparently
increased steady-state levels of ET-1 mRNA, prior to the major increase in cytoplasmic levels of
this mRNA, resulting from HIF-1α induction of the ET-1 gene. It remains to be seen whether this
55

is a general cytoplasmic mechanism for rapid induction of physiological modifiers preceding de
novo gene transcription needed for sustained expression.
In conclusion, our studies showed PlGF treatment of endothelial cells reduced levels of
miR-648, which binds the 3’UTR of ET-1 mRNA as a regulatory target. This post-
transcriptional mechanism affecting turnover of ET-1 mRNA, as illustrated in the regulatory
schematic (Fig. 8B), is proposed to be a means to negatively modulate ET-1 expression. The
miR-648 gene is located in the first intron of MICAL3 and our studies showed its expression was
co-transcriptionally regulated with MICAL3, by PAX5 from the distal promoter (P1; Fig. 6A).
Under basal conditions, levels of miR-648 are high, which maintain low levels of ET-1 mRNA
and protein; conversely in response to PlGF, the levels of miR-648 are reduced leading to
increased ET-1. The high levels of PlGF seen in SCD patients would have an expected outcome
of low PAX5 activity, as observed in vitro. This would result in reduced expression of miR-648
concomitantly leading to higher ET-1 levels (Fig. 8B). The net pathophysiological result would
be PHT as observed in SCA.
Ideally, further studies are warranted in an animal model, however human miR-648 does
not have a corresponding ortholog in mouse, thus this precludes studies in Berkeley sickle mice
or PlGF
-/-
mice (Sundaram et al., 2010). Our limited observations in SCA patients showed
significant differences in plasma levels of miR-648, compared to unaffected siblings as controls
(Fig. 8A). We feel further studies are needed in a larger SCA population to test the hypothesis
that miRNAs can be employed as prognostic biomarkers for pulmonary hypertension in SCA
patients. As a final thought, our work would suggest that an RNAi based therapeutic approach
may be beneficial in ameliorating PHT in SCA patients.
56

CHAPTER 4: Peroxisome Proliferator-activated Receptor α –mediated
Transcription of miR-199a2 Attenuates Endothelin-1Expression via Hypoxia-
inducible Factor-1α

4.1 Introduction
Endothelin and nitric oxide are two opposing vasoactive factors, which regulate vascular
tone. In blood vessels, the endothelin system plays an important role in basal vasoconstriction
and participates in the pathology of diseases such as hypertension, cardiovascular disease and
atherosclerosis (Kedzierski & Yanagisawa, 2001; Kisanuki et al., 2010). In the lungs, the
endothelin system controls the tonicity of both blood vessels and airways, and is involved in the
development of pulmonary hypertension (PH). PH occurs in 10-30% of patients with sickle cell
disease (SCD), and is associated with higher incidence of mortality in adults (Gladwin et al.,
2004; Klings, 2008; Minniti et al., 2009). A number of factors, such as chronic hemolysis,
hypoxia, hemostatic activation and inflammation have been implicated in endothelial
dysfunction, pulmonary vasoconstriction and remodeling in PH in SCD (Ataga et al., 2008;
Gladwin et al., 2004; Klings, 2008; Klings et al., 2014; Miller & Gladwin, 2012; Morris et al.,
2008; Zhang et al., 2014). Reduced bioavailability of nitric oxide due to quenching of NO by
extra-cellular hemoglobin (Bunn et al., 2010; Hsu et al., 2007; Nouraie et al., 2013; Wood et al.,
2013) and increased levels of endothelin-1 (ET-1) have been implicated in PH in SCD (Rybicki
& Benjamin, 1998; Sundaram et al., 2010).
The endothelin’s are a family of small peptides that include ET-1, ET-2 and ET-3, and
are produced primarily by vascular endothelial cells, and to a lesser extent by vascular smooth
muscle cells, airway epithelial cells, macrophages and fibroblast cells (Shao et al., 2011). ET-1
expression is induced by hypoxia subsequent to HIF-1α activation (Yamashita et al., 2001). We
57

showed that placenta growth factor (PlGF) induces the expression of ET-1 in cultured human
pulmonary microvascular endothelial cells via activation of hypoxia inducible factor (HIF-1α),
independently of hypoxia (Patel et al., 2008). Placenta growth factor is elaborated from bone
marrow erythroid cells and its levels are significantly increased in patients with chronic
hemolytic anemia, including SCD, due to a compensatory erythroid hyperplastic response
(Perelman et al., 2003). Moreover, our studies showed elevated PlGF levels in Berkeley sickle
(SS) mice and SCD patients are associated with increased levels of ET-1, and clinical markers of
pulmonary hypertension (Sundaram et al., 2010). These studies clearly indicate the importance
of PlGF in regulation of ET-1, and its role in PH in SCD.
PlGF induces HIF-1α activity (Patel et al., 2008; Nitin Patel et al., 2010), independently
of hypoxia. Others have shown participation of miR-199a as a post-transcriptional regulator of
HIF-1  under hypoxia for cell proliferation of non-small cell lung cancer cells (Ding et al.,
2013). However, the regulation of miR-199a availability in the context of SCD has not been
previously studied. This type of post-transcriptional regulation would be expected to directly
affect downstream HIF-1  -dependent gene expression events.
In the present study, we show that PlGF reduced miR-199a2 levels in cultured human
endothelial cells and that miR-199a2 targets the HIF-1α 3’UTR for turnover. The miR-199a2
precursor is transcribed as a Dynamin-3 opposite strand (DNM3os) RNA pol II transcript as
previously shown for the mouse DNM3 locus (Loebel, Tsoi, Wong, & Tam, 2005). The human
DNM3os transcript is also predicted to include miR-214 in the same transcription unit
(el Azzouzi et al., 2013). Our studies for the first time show that DNM3os and premir-199a2
transcription was co-regulated by PPARα. Moreover, PPARα agonist augmented the expression
of premir-199a2 and concomitantly attenuated the activation of HIF-1α, thus reducing the
58

expression of ET-1 in cultured endothelial cells. It is pertinent to note that Staels and coworkers
(Glineur et al., 2013) showed PPARα agonist, fenofibrate, attenuated ET-1 expression in
endothelial cells by increasing expression of transcriptional repressor Kruppel-like factor 11
(KLF11) and its binding to the ET-1 gene promoter. In our studies, fenofibrate acted to increase
transcription from the DNM3os locus leading to higher miR-199a2 expression with resultant
reduction of HIF-1  activity and subsequent reduction of ET-1 transcription.
These studies were corroborated in SCD patients, wherein we observed significantly
reduced plasma levels of miR-199a-5p, indicating that decreased levels of miR-199a-5p
contributed to increased levels of HIF-1α and its regulated gene, i.e. ET-1. We also showed that
feeding fenofibrate diet to wild type C57BL/6NJ mice and Berkeley sickle mice augmented
expression of miR-199a2, which resulted in reduced levels of ET-1 in lungs of both wild type
and sickle mice. These studies elucidate a novel molecular pathway linking up-regulation of
miR-199a2 to reduction of ET-1, thus providing a rationale for a therapeutic approach to
attenuate ET-1 levels and ameliorating pulmonary hypertension.

59

4.2 Materials and Methods
Reagents
Recombinant human PlGF was purchased from Peprotech (Rocky Hill, NJ). Clofibrate,
fenofibrate and GW 6471 were obtained from Sigma-Aldrich (St. Louis, MO). Primary
antibodies against PPARα, HIF-1α and secondary antibodies conjugated to horseradish
peroxidase (HRP) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Antisense
oligonucleotides (antagomirs) and miRNA-199a2 mimics were obtained from Shanghai
Genepharma (Shanghai, China) and their sequences are indicated in Table 1. The miR-199a2
over-expression vector was obtained from Genescript (Piscataway, NJ).
Human Subjects and Mice
All blood samples were obtained from children with homozygous SCA at steady state
(absence of sickle cell related acute event for three weeks prior to blood draw) at their elective
clinical visit for routine check-up and clinical blood draws. Blood samples were obtained with
the informed consent of the patient or parent/legal guardian using Institutional Review Board-
approved protocols through the Repository for Non-Malignant Hematology Specimens at
Cincinnati Children’s Hospital Medical Center. The plasma samples from the same SCD patients
(SS) and their unaffected sibling controls subjects were collected and stored at -80
o
C until
assayed. Animal protocols were approved by the Institutional Animal Care and Use Committee
at Cincinnati Children’s Hospital Medical Center. Berkeley sickle (BK-SS) mice originally
obtained from Jackson Laboratories were bred up to the 6
th
generation against a C57BL/6NJ
background. Berkeley sickle (BK-SS) mice carry deletions for the mouse α- and -globin genes,
and carry a transgene expressing the human -globin and a mini β-globin cluster including the β-
sickle globin (Pá szty et al., 1997; Nitin Patel et al., 2010). In–house bred C57BL/6NJ animals
60

served as controls. Six to ten animals per group were used for feeding. Five week old
C57BL/6NJ and BK-SS mice were fed with either the Mod Test Diet 59M3 (referred to as sickle
cell mouse diet) from Purina (Richmond, Indiana) or same diet supplemented with 0.05%
(wt/wt) of fenofibrate for 10 weeks. At the end of the study mice were bled and lungs were
removed, and stored at -80
o
C for assay of miRs and ET-1.
Endothelial cell culture
The immortalized human dermal microvascular endothelial cells, HMEC-1, originally
developed by Drs Edwin Ades and Francisco J. Candall of the Centers for Disease Control and
Prevention (CDC), and Dr. Thomas Lawley of Emory University, were obtained from the CDC
(Atlanta, GA). HMEC-1 cells were cultured in RPMI-1640 media containing 10% FBS, heparin,
endothelial cell mitogen and MEM vitamins as previously described (K. S. Kim et al., 2006).
Human lung microvascular endothelial cells (HLMVEC), primary cell cultures at passage two,
were obtained from Cell Applications (San Diego, CA). HLMVEC were grown in microvascular
endothelial cell media (Cell Applications, San Diego, CA) according to vendor’s instructions.
These cells displayed an endothelial cell phenotype and were positive for PECAM-1 (CD31) and
von Willebrand factor. HLMVEC were used up to passage 8-9, without undue effect on
morphology or experimental outcome. Unless otherwise specified, HLMVEC and HMEC were
kept overnight in complete media (without growth factors) containing 2% FBS, followed by
serum- free media for 3 hrs, before treatment with either PlGF (250 ng/ml) or other experimental
conditions (Patel, Gonsalves, Yang, Malik, & Kalra, 2009b).
RNA interference
The shRNA clones were a generous gift from Dr. Jae Jung (USC). The human shRNA
library was purchased from Open Biosystems (now Thermo Scientific, Grand Island, NY). All
61

synthetic shRNA sequences were supplied in a pGIPZ lentivector. The PPAR  shRNA (clone #
0197-0066-G8) sequence is as follows:
TGCTGTTGACAGTGAGCGCCGATCAAGTGACATTGCTAAATAGTGAAGCCACAGAT
GTATTTAGCAATGTCACTTGATCGTTGCCTACTGCCTCGGA.
Transient Transfections
1 x 10
6
HMEC-1 cells were transfected with shRNA plasmid (1 µ g), luciferase reporter
plasmid (1 µ g), either miR-199a2 mimics or inhibitors (90 pmoles) by nucleofection, utilizing
RPMI-1640 as the transfection solution and program Y-001 on the Amaxa Nucleofector (Lonza,
Allendale, NJ). The transfection was highly effective in the absence of nucleofection agent (J.
Kang et al., 2009). Transfected cells were transferred to complete media and incubated
overnight. Cells were kept in serum free media for 3 hours prior to treatment with PlGF (250
ng/ml).
Luciferase Reporter vectors
A 2- Kb segment containing the DNM3os promoter was amplified by PCR, utilizing a
BAC clone as the template (BACPAC Resource Center, Oakland, CA; Clone # RP11-455 O13),
and the resulting PCR product was ligated into pGL3-Basic luciferase reporter vector as per
manufacturer’s instructions (Promega, Madison, WI). The subcloned wild-type 2 Kb DNM3os
promoter was used as a template for generating mutations at specific sites. The mutations in three
PPARα sites were generated by the Q5 Site-Directed mutagenesis kit (New England Biolabs,
Ipswich, MA) utilizing primers listed in Table 1. The three PPAR  mutations are designated as
PPmt1 (bases -216/-206), PPmt2 (-317/-307) and PPmt3 (-789/-779). Double mutations in the
PPmt1 and PPmt2 sites were generated using the PPmt1 construct as a template. The 684 bp
HIF-1  3'UTR luciferase reporter construct (3’UTR positions +1 to +684) was cloned using
62

primers listed in Table 1, and the PCR product was ligated into the XbaI cut pGL3-Control
vector. Mutations within the miR-199a2 binding sites in the HIF-1  3'UTR were generated using
the Q5 Site-directed mutagenesis kit and primers listed in Table 1. All constructs and sequence
mutations were verified by DNA sequencing (Retrogen, San Diego, CA).
Isolation of RNA and quantitative PCR
Total RNA was isolated from cells using TRizol reagent (Life Technologies, Grand
Island, NY). mRNA expression was quantified using specific primers (Table 2) by quantitative
real time PCR (qRT-PCR). qRT-PCR of mRNA templates (100 ng) were performed using the
One-Step SYBR PrimeScript Kit (Clontech Laboratories, Mountain View, CA) and the ABI
Prism 7900HT sequence detection system (Applied Biosystems, Foster City, CA). Amplification
was carried out as follows: Step 1: Reverse Transcription, 42º C for 5 min, 95º C for 10 s, Step 2:
PCR reaction (40 cycles), 95º C for 5 s, 60º C for 34 s, followed by detection. Values are
expressed as relative levels of RNA normalized to the mRNA levels of the housekeeping gene
GAPDH. Relative quantification of the mRNA levels were calculated using 2
-ΔΔCt
by the
comparative threshold cycle method (Ct), where ΔΔCt= (Ct of target gene of treated sample - Ct
of GAPDH mRNA of treated sample) - (Ct of target gene of untreated sample - Ct of GAPDH
mRNA of untreated sample)(Patel et al., 2009b).
63

Table 2. Oligonucleotides primers used in the Chapter 4 study.
64

Isolation and quantification of miRNAs
microRNAs were purified utilizing the mirVana isolation kit (Life Technologies, Grand
Island, NY) according to the manufacturer's protocol. miRNA expression levels were determined
by using the TaqMan MicroRNA assay kits for indicated miRNA (Invitrogen, Carlsbad, CA),
according to the manufacturer's instructions (Patel, Tahara, Malik, & Kalra, 2011b). miRNA
expression was quantified by qRT-PCR and expression levels were normalized to a reference
gene, RNU6B (U6 snRNA) (Patel et al., 2011b) using the comparative threshold cycle method
(Ct). miRNAs were also quantified by Northern blot analysis. Briefly, 35 µ g of total RNA was
run on a 15% non-denaturing polyacrylamide gel. The RNA was transferred to a Biodyne B
nylon membrane. The membrane was cross-linked under UV light and pre-hybridized for 30 min
using the UltraHyb hybridization buffer (Ambion, Grand Island, NY) at 42º C. The membrane
was then hybridized with biotin-labeled probes for miR-199a2 and 5S rRNA, synthesized at
Valugene (San Diego, CA), at 65º C overnight. The membrane was washed twice in washing
buffer (Thermoscientific, Rockford, IL), followed by blocking with 5% non-fat milk in 1XPBS
at room temperature. Streptavidin-HRP (1:250 dilution) was added to the membrane, incubated
at room temperature for 3 hr, and followed by 2 washes with washing buffer according to
vendor’s instructions. The membranes were developed utilizing Clarity Western ECL substrate
(BioRad, Richmond, CA), and resulting images quantified using the Image J analysis software.
Western blot analysis
Cells were lysed in RIPA buffer and 25 µ g of protein extracts were subjected to SDS-
PAGE, followed by electrophoretic transfer, Western blotting and chemiluminescence detection
of bound antibody, as previously described (Giri, Selvaraj, & Kalra, 2003). Membranes were
65

probed with indicated dilutions of anti-HIF-1α (1:250, sc-12542, Santa Cruz Biotechnology,
Santa Cruz, CA), anti-PPARα (1:250, sc-9000, Santa Cruz Biotechnology), anti-ET-1 (1:250,
Santa Cruz Biotechnology), and HRP- conjugated anti-goat or anti-rabbit secondary antibodies
(1:10,000, Sigma Aldrich, St. Louis, MO). Membranes were stripped and re-probed with anti-β-
actin (1:10,000, Sigma Aldrich). Quantitative analysis was performed using the Image J analysis
software.
Quantification of secreted ET-1
HMEC-1 (1x10
5
cells) were transfected with indicated miRNA or sc-miRNA (90 pmole)
in complete media and incubated at 37° C for 24 hr. Cells were washed with serum-free medium
(SFM) and incubated for 3 h in SFM (2 ml). Treatments were begun by replacement of SFM
with fresh SFM (1 ml). Cells were treated with either PlGF (250 ng/ml) or fenofibrate (0.1 mM)
overnight. The culture supernatant was collected and an aliquot (0.1 ml) was assayed for ET-1
release using an enzyme-linked immunosorbent assay (ELISA) kit (Assay Design/Enzo Life
Sciences, Farmingdale, NY) (Patel et al., 2008). The cells were isolated by scraping and the
pellet was assayed for protein content utilizing the Bradford method. Plasma specimen ET-1
levels were determined using DuoSet ELISA kit from R and D (Minneapolis, MN)
Chromatin immunoprecipitation (ChIP) assay
HMEC-1 (5x10
6
cells) cultured in 150 mm petri dishes were kept in SFM for 3 h and
treated with fenofibrate (0.1 mM) or fenofibrate + GW6471 (5M) for 2 hr. ChIP analysis was
performed using antibody to PPARα (Santa Cruz Biotechnology, Santa Cruz, CA) (Patel et al.,
2009b). Briefly, immunoprecipitated DNA was air-dried and resuspended in nuclease free water.
DNA samples were subjected to PCR amplification utilizing primers for the DNM3os promoter
regions of interest corresponding to PPARα binding sites (Table 2). PCR was performed for 35
66

cycles under the following conditions: denaturation at 95° C for 30 s, annealing at 55° C for 60 s,
and extension at 72° C for 2 min. The PCR products were subjected to electrophoresis on a 2%
agarose gel, visualized by ethidium bromide staining, and quantified using the Image J analysis
software.
Statistical analysis
The significance between two groups was ascertained using an unpaired Student’s t-test,
and results are presented as means ± S.E.M. *P<0.05, **P<0.01, ***P <0.001, and ns (not
significant) with P>0.05.

67

4.3 Results
Interaction of miR-199a-5p with the 3’-UTR of HIF-1α mRNA modulates HIF-1α mRNA
levels
Previous studies from our lab show PlGF induces HIF-1α in HMEC-1, independent of
hypoxia (Patel et al., 2008). Moreover, as previously described in Chapter 3, the expression of
miR-199a-5p was reduced by ~10-fold, in response to PlGF. A predicted miRNA recognition
element (MRE) for miR-199a-5p is located within nucleotides +16 to +38 of the 3’-UTR of HIF-
1α mRNA (Fig. 12A). For these reasons, we examined the binding of miR-199a-5p to its target
sequence in a luciferase reporter (pGL3-HIF-1 -3’UTR) in response to PlGF (Fig. 12B). We co-
transfected pGL3-HIF-1α-3’UTR with exogenous miR-199a-5p mimic, we observed ~50%
decrease in luciferase activity, compared to the luciferase reporter itself (Fig. 12C, lane 2 vs. lane
1). Co-transfection of the same reporter with anti-miR-199a-5p quenched endogenous miR-199a-
5p showed a ~90% increase in luciferase activity (Fig. 12C, lane 3 vs. lane 1). Further,
scrambled (sc) mimic or sc inhibitor were used as negative controls, which did not change
luciferase activity significantly (Fig. 12C, lanes 4 and 5 vs. lane 1).
68

Figure 12. PlGF attenuates miR-199a2-5p expression, which targets the 3’UTR of HIF-1α mRNA. A.
Schematic representation of miR-199a2-5p location in 3’UTR of HIF-1α mRNA. B. Schematic
representation of the HIF-1α -3’-UTR reporter plasmid. The region between bases +1 to +684 of the HIF-
1α 3’UTR containing the predicted target site for miR-199a2 was inserted at the 3’-end of luciferase
reporter gene in pGL3-Control plasmid. C and D. HMEC were transfected with HIF-1α -3’UTR reporter
plasmid with indicated miR- mimic, anti-miR, sc miR (NC, negative control) or sc-anti-miR (NC). E.
Schematic of mutation in miR-199a2 MRE in 3’UTR of HIF-1α mRNA. Asterisks indicate mutated bases. F.
HMEC were transfected with wild-type HIF-1α -3’UTR plasmid or the indicated mutant HIF-1α 3’UTR
plasmid, with indicated miR or anti-miR.

69

Since PlGF decreased endogenous miR-199a-5p levels, exogenous miR-199a-5p under
these conditions would be expected to attenuate the expected increase of HIF-1α mRNA
stability. Indeed, co-transfection of pGL3-HIF-1α-3’UTR with miR-199a-5p followed by PlGF
treatment resulted in ~100% decrease in luciferase reporter activity (Fig. 12D, lane 3 vs. lane 2).
The primary effect was attributed to the exogenous miR-199a-5p because transfection with anti-
miR-199a-5p alone had no effect on luciferase activity consistent with down- regulation of
endogenous miR-199a in PlGF treated cells (Fig. 12D, lane 4 vs. lane 2). Cells transfected with
scrambled (sc) mimic or inhibitor did not affect reporter activity compared to PlGF-treated cells
(Fig. 12D, lanes 5 and 6 vs. lane 2). Next, we mutated the miR-199a-5p seed sequence
complement in pGL3-HIF-1α-3’UTR as indicated in the schematic (Fig. 12E). As shown in
Fig.12F, mutation of the miR-199a-5p binding site reversed the reduction of reporter luciferase
activity under basal conditions (Fig. 12F, lane 2 vs. lane 1). The HIF-1α 3’-UTR mt, which
depleted the miR-199a-5p binding site did not show any change in luciferase activity upon co-
transfection with either miR-199a-5p or anti-miR-199a-5p (Fig. 12F, lane 3 vs. lane 2, and lane 4
vs. lane 2). Taken together, these data are consistent with miR-199a-5p binding to its predicted
target site in the 3’UTR of HIF-1α mRNA for the purpose of modulating HIF-1α synthesis via
mRNA turnover.

miR-199a-5p modulates ET-1 mRNA expression via its effect on the levels of HIF-1α
Since our previous results showed that miR-199a-5p was directly binding to the 3’UTR
of HIF-1α mRNA, we examined the effects of exogenous miR-199a-5p and anti-miR-199a-5p on
endogenous HIF-1α mRNA levels. Transfection with miR-199a-5p mimic reduced basal HIF-1α
mRNA by ~75%, and anti-miR-199a-5p effectively antagonized endogenous miR-199a-5p,
70

resulting in ~2-fold increase in HIF-1α mRNA (Fig. 13A). Both negative controls, sc miR mimic
and sc miR inhibitor did not affect basal HIF-1α mRNA expression (Fig. 13A).
Previous study from our lab showed PlGF induces ET-1 mRNA and protein expression via
activation of HIF-1α, independently of hypoxia (Patel et al., 2008). Since miR-199a-5p affected
HIF-1α mRNA levels, we examined whether HIF-1α dependent ET-1 expression was negatively
affected as well. Transfection of miR-199a-5p mimic into HMEC-1 indirectly attenuated basal
levels of ET-1 mRNA (Fig. 13B, lane 2 vs. lane 1) while anti-miR-199a-5p augmented by ~1.6-
fold ET-1 mRNA expression (Fig. 13B, lane 3 vs. lane 1) under basal conditions. The 3’UTR of
ET-1 mRNA has no miR-199a-5p binding site(s), thus these data are consistent with miR-199a-
5p indirectly affecting ET-1 mRNA expression by reducing HIF-1α synthesis at the level of
mRNA translation.
71

72

Effect of miR-199a-5p mimics and anti-miR-199a-5p on PlGF-mediated expression of HIF-
1α and ET-1 mRNAs
Since PlGF mediated HIF-1α expression augments ET-1 expression (Patel et al., 2008),
we examined the role of miR-199a-5p in the expression of HIF-1α and ET-1 mRNA. As shown
in Fig. 13C, transfection of miR-199a-5p mimic in HMEC followed by PlGF treatment resulted
in ~65% reduction in HIF-1α mRNA level compared to PlGF treated cells (Fig. 13C, lane 3 vs.
lane 2). Conversely, transfection with anti-miR-199a-5p restored HIF-1α mRNA expression over
and above that seen in PlGF treated cells (Fig. 13C, lane 4 vs. lane 2). The effects of scrambled
(sc) mimic and sc inhibitor were not significant (Fig. 13C, lanes 5 and 6 vs. lane 2). Transfection
of miR-199a-5p mimic in the presence of PlGF attenuated ET-1 mRNA expression by~75% as
expected for miR-199a-5p directed reduction of HIF-1α expression (Fig. 13D, lane 3 vs. lane 2).
Conversely, anti-miR-199a-5p did not significantly alter ET-1 mRNA levels compared to PlGF-
treatment alone (Fig. 13D, lane 4 vs. lane 2). Both negative controls, sc miR and sc anti-miR,
had no effect on PlGF-mediated ET-1 mRNA expression (Fig. 13D, lanes 5 and 6 vs. lane 2).
Moreover, PlGF mediated HIF-1α protein expression was completely attenuated by miR-199a-5p
mimic (Fig. 13E, lane 3 vs. lane 2), while negative control, sc miR mimic had modest or no
effect on HIF-1α protein levels (Fig. 13E, lane 4 vs. lane 2). Additionally, PlGF-mediated ET-1
Figure 13. miR-199a2 targets the 3’UTR of HIF-1α mRNA affecting expression of downstream target
gene ET-1. HMEC were transfected with indicated miR, anti-miR and sc-miR and sc-anti-miR. 24 h post-
transfection, cells were treated with PlGF for 6 h where indicated. RNA and protein were isolated from
separately treated cells. mRNA was quantified by qRT-PCR (A-D) and western blots were performed for
HIF-1α (panel E) and ET-1 (panel F). β-actin immunodetection was used for normalization of protein
loading. G and H, Effect of transfection of miR-199a-5p mimic and anti-miR-199a on levels of miR-199a in
HMEC. HMEC were transfected with miR-199 mimic, anti-miR-199 or sc mimic, at 90 pmol, for 24 h,
followed by extraction and quantitation of miR-199a by Northern blotting.
73

protein expression was reduced by miR-199a-5p mimic (Fig. 13F, lane 3 vs. lane 2), while the
effect of sc miR mimic was modest (Fig. 13F, lane 4 vs. lane 2). Furthermore, changes in
intracellular levels of miR-199a in HMEC transfected with miR-199a-5p mimic and antimir-
199a were confirmed by Northern blot (Fig. 13G and H). Taken together, the data showed miR-
199a-5p mimic attenuated PlGF-mediated expression of both HIF-1α mRNA and protein, and
subsequently caused reduction of ET-1 mRNA and protein expression.

PlGF attenuates expression of DNM3os, premir-199a2 and premir-214 mRNA
Previous studies (Y.-B. Lee et al., 2009) show that murine miR-199a2/miR-214 cluster is
located in the anti-sense strand of Dynamin 3 (DNM3), referred to as Dynamin 3 opposite strand
(DNM3os). Human DNM3os gene also produces a transcript that includes premir-199a2 and
premir-214 (Fig. 14A). Furthermore, in silico analysis revealed that mature miR-199a-5p and
miR-199a-3p also originate from premir-199A1 as an opposite strand transcript arising from an
intron of DNM2 (Fig. 14A). The miRNAs encoded by miR-199B are similar in nucleotide
sequence to those encoded by miR-199A1 and miR-199A2, but have different seed sequences
(Fig. 14A). By contrast the latter two miRNA genes produce identical, mature miRNAs, hence
the name miR-199a. In order to establish the origin of miR-199a-5p, it became necessary to
examine the pattern of expression for premir-199a1, premir-199a2 and premir-199b, in response
to PlGF.
Following PlGF treatment of HMEC, we observed ~50% reduction of premir-199a2 with no
change in premir-199a1 expression (Fig. 14B). In contrast, premir-199b expression increased by
~2-fold (Fig. 14B). These data showed PlGF selectively reduced expression of premir-199a2,
which we concluded was the precursor of mature miR-199a-5p under these conditions. Next, we
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examined whether premir-199a2-5p and premir-214, also located in DNM3os, were co-expressed
during DNM3os transcription. Treatment of HMEC with PlGF for 6 h showed ~60% decrease in
the expression of DNM3os RNA (Fig. 14C) and a concomitant decrease in the expression of
premir-199a2 (Fig. 14B) and premir-214 (Fig. 14C).
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Figure 14. PlGF attenuates expression of miR-199a2 and its host gene DNM3os in endothelial cells.
A. Summary of premirs and miRNA sequences originating from DNM1os, DNM2os and DNM3os loci,
respectively. B and C. HMEC were treated with PlGF for 2 hr, and expression of indicated premirs
determined in extracted RNA by qRT-PCR. D. HMECs were exposed to hypoxia (1% O
2
) for 24 hr,
followed by extraction of RNA for qRT-PCR analysis of indicated pre-miRs and DNM3os.
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HIF-1α is activated by different types of stress (Pereira, Frudd, Awad, & Hendershot,
2014; Semenza, 2012). In an effort to determine whether other activators of HIF-1α were
operative in transcriptional regulation of DNM3os we asked whether hypoxia has such an effect.
Hypoxic stress stabilizes HIF-1α protein, thus we examined whether this stimulus also
modulated expression of DNM3os RNA, premir-199a2 and premir-214. Exposure of HMEC to
hypoxia (1% O
2
) for 16 h reduced the expression of DNM3os RNA, premir-199a2 and premir-
214 (Fig. 14D). These results showed that both PlGF-mediated signaling and hypoxia attenuated
transcription of DNM3os, premir-199a2 and premir-214 in a coordinated manner.

DNM3os transcription is regulated through PPARα
Bioinformatic analysis of the ~2 kb 5’-flanking region of human DNM3os (NCBI Gene
ID 100628315) revealed the presence of three PPARα binding sites, relative to the transcription
start site, as shown in the promoter schematic (Fig. 15A). The involvement of PPARα in
DNM3os transcription was established by transfection of HMEC-1 with PPARα shRNA, which
resulted in ~80% reduction of overall DNM3os transcription and concomitant ~90% reduction in
premiR-199a2 and ~70% reduction in premiR-214 (Fig. 15B). Furthermore, analysis of these
cells for expression of PPARα showed that shRNA treatment specifically reduced PPARα
mRNA and PPARα protein as shown in Figs. 15C and 15D, respectively, indicating that
PPARα
participation in basal transcription of DNM3os, premiR-199a2 and premiR-214.
A secondary approach was employed to establish the participation of PPARα in DNM3os
transcription. PPARα activity can be modulated by pharmacological agonists such as fenofibrate
and clofibrate (Gervois, Fruchart, & Staels, 2007). Treatment of HMEC-1 with fenofibrate
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showed a time dependent increase in PPARα mRNA levels, which peaked between 4-8 h (data
not shown). Fenofibrate treatment of HMEC-1 for 4 h, in the absence of PlGF, resulted in ~4-
fold increase in PPARα mRNA level (Fig. 15E, lane 2 vs. lane 1). Treatment of HMEC-1 cells
with the antagonist GW6471 inhibited fenofibrate induced PPARα mRNA expression (Fig. 15E,
lane 3 vs. lane 2). Treatment of HMEC-1 with clofibrate, another PPARα agonist, showed a 2.5-
fold increase in PPARα mRNA (Fig. 15E, lane 4 vs. lane 1), which was attenuated to the basal
level by GW6471 (Fig. 15E, lane 5 vs. lane4). The actions of PPARα agonists and antagonists
were also reflected in the corresponding PPARα protein levels (Fig. 15F). Fenofibrate treatment
of HMEC-1, increased DNM3os RNA expression by ~4.5-fold (Fig. 15G lane 2 vs. lane 1),
which was attenuated by ~90% in the presence of GW6471 (Fig. 15G. lane 3 vs. lane 2).
Furthermore, clofibrate increased DNM3os transcription by ~4-fold, which was reduced by
~70% in the presence of GW6471 (Fig. 15G, lane 5 vs. lane 4). Taken together, these results
showed that PPARα was involved in DNM3os transcription in HMEC-1.
Since miR-199a2 and miR-214 are processed from the DNM3os transcript, we examined
whether PPARα agonists similarly affected transcription of premiR-199a2 and premiR-214. As
shown in Fig. 15G, fenofibrate treatment resulted in augmented expression of premiR-199a2 by
~3-fold, while PPARα antagonist GW6471 attenuated premir-199a2 levels to the basal level
(Fig. 15G, lane 3 vs. lane 2). Furthermore, clofibrate increased premiR-199a2 mRNA expression
by ~3.5-fold compared to untreated sample (Fig. 15G, lane 4 vs. lane 1), while GW6471
attenuated premiR-199a2 expression (Fig. 15G, lane 5 vs. lane 4). Since premir-214 is part of the
miR-199a2 cluster, we examined its expression in response to PPARα agonist. Both fenofibrate
and clofibrate augmented premir-214 expression while GW6471 antagonist inhibited premir-214
expression (Fig. 15G). Furthermore, mature miR-199a2 expression increased ~3.5-fold in
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response to fenofibrate, which was attenuated by GW6471 (Fig. 15H). Northern blotting showed
PlGF reduced the expression of mature miR-199a, while fenofibrate augmented miR-199a
expression (Fig. 15I), thus confirming the results obtained by qRT-PCR. The effect of PPARα
agonist on DNM3os transcription was specific as troglitazone, a PPARγ agonist, was ineffective
in augmenting DNM3os transcription (Fig. 15J). Taken together these data showed that miR-
199a2 and miR-214 co-located in DNM3os were indeed co-transcriptionally regulated by
PPARα.
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Involvement of PPARα cis- binding elements in DNM3os transcription
Since PPARα agonists augmented DNM3os transcription, we needed to determine
whether one or more of the putative PPAR  sites were required for DNM3os transcription. For
this analysis, a transcription reporter was constructed from 2 kb of the 5’ flanking region
immediately upstream of DNM3os fused to the luciferase coding region in pGL3-Basic, shown
in schematic form in Fig. 16A. Transfection of HMEC-1 with the reporter driven by the wild
type (wt) DNM3os promoter (pDNM3os-luc) followed by treatment with fenofibrate showed
Figure 15. PPARα regulates the transcription of miR-199a2 and its host gene DNM3os. A. Schematic
representation of PPARα cis-binding elements in promoter of DNM3os. B-F. HMECs were either
transfected with PPARα shRNA (24 h) or treated with indicated agents for 4 h. RNA and protein were
separately isolated from parallel cultures. RNA was subjected to qRT-PCR for indicated premirs,
DNM3os, PPARα and mature miR199a2. B Effect of transfected PPARα shRNA on expression of
PPARα mRNA (B) and C. PPARα protein. D. Effect of transfection of HMEC with PPARα shRNA on
expression of DNM3os, premir-199a2 and premir-214. E. Effect of PPARα agonist and antagonist on
with antibody to PPARα and β-actin was used as a loading control. Western blot are representative of
three independent experiments. G. Effect of PPARα agonist and antagonist on RNA levels of DNM3os,
premir-199a2 and premir-214. H. Effect of PPARα agonist and antagonist on mature miR-199a-5p
levels in HMEC. I. Northern blot for miR-199a2 expression in HMEC treated with either PlGF or
fenofibrate. J. Effect of PPARα and PPARα agonist on DNM3os RNA levels. Data are representative of
three independent experiments.
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~1.8-fold increase in luciferase activity compared to untreated cells (Fig. 16B, lane 2 vs. lane 1).
By contrast, co-treatment with fenofibrate and GW6471 (antagonist) reduced luciferase activity
to the background level (Fig. 16B, lane 3 vs. lane 2), confirming the role of PPARα. Moreover,
clofibrate augmented reporter expression by ~1.6-fold (Fig. 16B, lane 4 vs. lane1) and GW6471
prevented induction beyond the background level of luciferase expression (Fig. 16B, lane 5 vs.
lane 4). Taken together these data showed that PPARα agonists indeed activated the DNM3os
promoter and PPARα antagonist inhibited DNM3os induction.
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The wt promoter was mutated at each of the predicted PPARα binding sites (Figs. 15A
and 16A) by replacing the wt core sequences with mutated sequences as indicated in Fig. 16A.
Transfection and assay of single PPARα site mutants (sites 1 and 2) of the DNM3os promoter
resulted in unchanged luciferase activity in response to fenofibrate as compared to wild type
DNM3os promoter (Fig. 16C). Similarly, the double mutation of sites 1 and 2 was unresponsive
to fenofibrate (Fig. 16C). By contrast, PPARα site 3 mutant remained responsive to fenofibrate
(Fig. 16C) indicating that this site was non-essential for PPARα-mediated trans-activation. These
results showed that the proximal PPARα cis-binding elements (sites 1 and 2) but not site 3
present in the DNM3os promoter were essential for fenofibrate mediated transcription of
DNM3os.
Finally, we determined whether PPARα binds to the DNM3os promoter in native
chromatin of HMEC by performing chromatin immuno-precipitation (ChIP) analysis. For this
analysis chromatin was prepared from untreated, fenofibrate treated, and fenofibrate plus
GW6471 treated HMEC-1 cells. Chromatin preparations were immunoprecipitated with either
antibody to PPARα or control non-specific rabbit IgG. The PCR products corresponding to each
PPARα site in the DNM3os promoter, amplified with appropriate primers (listed in Table 1),
were detected and quantitated following gel electrophoresis. HMEC-1 cells treated with
Figure 16. PPARα cis-binding elements in promoter ofDNM3osregulate expression ofDNM3os as
determined by reporter assay and ChIP. A, schematic of mutations in core sequences corresponding to
PPARα sites (1, 2, and 3; see Fig. 4A) in DNM3os promoter reporter constructs. B and C, HMEC cells
were transfected with either DNM3os-luc (WT) plasmid or the mutant plasmid with singly mutated
PPARα binding sites (1, 2, or 3) or double mutation of PPARα sites1 and 2 in DNM3os-luc constructs
(PPmt). Transfected cells were treated as indicated with PPARα agonists (fenofibrate or clofibrate) or
antagonist (GW6471) for 24 h. The cell lysates were assayed for luciferase activity and normalized to
Renilla activity. D –F, HMEC cells were treated with either fenofibrate or fenofibrate+GW6471 for 2 h.
The soluble chromatin was isolated and immune-precipitated with either antibody to PPARα or control
rabbit IgG.
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fenofibrate showed a ~1.5 fold increase in the expected 111-bp PCR product (corresponding to
amplicon 1), a ~1.4 fold increase in expected 137 bp PCR product (corresponding to amplicon 2)
and no change in expected 113 bp PCR product (corresponding to amplicon 3) as shown in Fig.
16 D, E and F. The augmented binding of PPARα to the promoter region was completely
reversed when cells were treated with PPARα antagonist (GW6471) as shown in Fig. 5D and 5E.
The ChIP analysis was specific since PCR products were not detected in the control IgG
immunoprecipitates (Fig. 5 D, E and F, bottom panel) and furthermore the input DNA used for
PCR amplification was equivalent in all samples (middle panels in Figs. 5 D, E and F). Together
with results above, these data showed that fenofibrate augmented binding of PPARα to proximal
sites 1 and 2 of the DNM3os promoter leading to increased transcription of DNM3os in vitro and
in vivo.

PPARα agonists mediate expression of miR-199a-5p, in turn regulating expression of HIF-
1α and a target gene ET-1
Since we showed that PPARα agonist increased levels of miR-199a2-5p, which targets
HIF-1α mRNA, we anticipated that fenofibrate would reduce HIF-1α activity and subsequently
down regulate expression of its downstream target gene ET-1. Indeed, treatment with fenofibrate
reduced HIF-1α mRNA levels by ~80% (Fig. 6A, lane 2 vs. lane 1), while co-treatment with
GW6471 restored HIF-1α mRNA expression (Fig. 6A, lane 3 vs. lane 2). Similar results were
obtained with clofibrate, which showed ~80% reduction in HIF-1α mRNA expression (Fig. 6A,
lane 4 vs. lane 1). The effect of PPAR  agonist was reversed in the presence of GW6471
(Fig.7A, lane 5. vs lane 4). These results showed that PPARα agonists attenuated HIF-1α levels
by mRNA turnover or translation inhibition following increased miR-199a2-5p expression.
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Next, we examined the effect of PPARα agonists on ET-1, a downstream gene regulated by
HIF-1  and a key mediator of PHT in SCA (Sundaram et al., 2010). As expected, ET-1 mRNA
levels were reduced by ~80% and ~60%, by fenofibrate and clofibrate respectively, in HMEC-1
(Fig. 6B lanes 2 and 4 vs. lane 1); moreover, GW6471 reversed the effects of PPARα agonists on
ET-1 mRNA expression (Fig. 6 B, lane 3 vs. lane 2 and lane 5 vs. lane 4). In support of a
mechanistic model involving premir-199a2/miR-199a2 synthesis, fenofibrate treatment of
HMEC resulted in a decrease in HIF-1α protein levels by ~70% (Fig. 6C) and a corresponding
reduction of ET-1 protein by ~90% (Fig. 6D). As a control for PPARα activation, the GW6471
antagonist reversed the observed effects of fenofibrate on HIF-1α (Fig. 6C) and ET-1 proteins
(Fig. 6D).
In order to confirm the physiological relationship of PPARα to HIF-1α and ET-1, we
examined whether the PPARα agonists affected primary human endothelial cells similarly to the
effects observed in HMEC-1. Treatment of primary human lung microvascular endothelial cells
(HLMVEC) with fenofibrate resulted in ~75% reduction of HIF-1α mRNA levels (Fig. 6E, lane
2) and ~60% reduction of ET-1 mRNA levels (Fig. 6E, lane 2). In addition, GW6471 reversed
the effect of fenofibrate on the expression of HIF-1α mRNA (Fig. 6E, lane 3 vs. lane 2) and ET-1
mRNA (Fig. 6E, lane 3 vs. lane 2).
Our previous studies show PlGF augments the release of ET-1 from HPMVEC. Herein, we
examined whether PlGF mediated ET-1 release was modulated by miR-199a mimic in HMEC-1.
As shown in Fig. 6F, PlGF caused ~75% increase in ET-1 release, which was attenuated by
~65% below basal expression with miR-199a mimic but not with sc miR mimic. Similarly,
PlGF-induced ET-1 release was 100% inhibited by fenofibrate (Fig. 6F, lane 6 vs. lane 2). These
results showed that PlGF-induced secretion of ET-1 from HMEC was attenuated by miR-199a2.
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Furthermore, these results showed that PPARα agonist reduced expression of HIF-1α and its
downstream target gene ET-1 by up-regulating the expression of miR-199a2, which is
responsible for post-transcriptional regulation of HIF-1 .
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Figure 17. PPARα agonists attenuate expression of HIF-1α and ET-1 in both HMEC cells and primary
HLMVEC. A and B, qRT-PCR was performed for HIF-1α and ET-1 mRNAs. C and D, Western blot was
performed on protein extracts utilizing antibody to HIF-1α, ET-1, and β-actin (loading control). The data
are representative of three independent experiments. E, HIF-1α and ET-1 expression in HLMVEC
treated with fenofibrate or fenofibrate+GW 6471.

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Does PlGF mediated induction of ET-1 expression involves Kruppel-like factor11 and HIF-
1?
Recent studies show fenofibrate attenuates ET-1 expression by a PPAR-dependent
mechanism via induction of Kruppel like factor 11 repressor, and by a PPAR-independent
mechanism requiring inhibition of glycogen synthase kinase-3 activity. In the present study, we
examined the role of KLF-11 and HIF-1 in the transcriptional regulation of ET-1 expression, in
response to PlGF and fenofibrate.
We examined the activity of the full length ET-1 promoter (669 bp) (Hu et al., 1998;
Patel et al., 2008), which has three KLF-11 cis-binding sites and a single HRE site, with a
luciferase reporter gene as depicted in Fig. 17G. Treatment of HMEC-1 with PlGF increased the
wt ET-1 promoter (669 bp) activity by ~2-fold (Fig. 17 H, lane 2 vs. lane 1), while fenofibrate
reduced PlGF induction of luciferase expression to the basal level (Fig. 17H, lane 3 vs. lane 2).
Next, we examined the activity of a truncated ET-1 promoter (176 bp) in the context of the
luciferase reporter. Note that the truncated promoter completely lacks the three KLF-11 sites (Hu
et al., 1998), (see Fig. 17G). HMEC transfected with truncated ET-1 promoter showed ~1.4-fold
increase in basal luciferase activity compared to wt ET-1 promoter (669bp), in the absence of
PlGF (Fig. 17H, lane 4 vs. lane 1). This indicated to us that deletion of the KLF-11 sites removed
any repressive activity of KLF-11 on reporter activity.
In response to PlGF treatment, the activity from the truncated promoter was substantially
the same as observed with wt ET-1promoter (Fig. 17H, lanes 5 vs. lane 2), indicating that
deletion of KLF-11 site in the truncated ET-1 promoter did not change maximum reporter
activity. In an effort to determine the relative contribution of KLF-11 and HIF-1  on ET-1
induction, we noted that fenofibrate treatment with the truncated ET-1 promoter reduced
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transcription activity to the level observed in the absence of PlGF (Fig. 17H, lane 6 vs. lane 4).
We interpreted this as a ~60% reduction of induction, accounting for the difference in basal
activities exhibited by the wt and truncated promoters (Fig. 17H, lane 4 vs. lane 1). The
requirement for HIF-1  in ET-1 transcription was demonstrated with the truncated ET-1
promoter reporter, where both fenofibrate and exogenous miR-199a mimic reduced ET-1
promoter activity due to reduction of HIF-1  activity (Fig. 17H, lanes 6 and 7, respectively).
By a similar argument, the contribution of KLF-11 to PlGF induction of ET-1
transcription was ~40% based on the observed increase in basal activity of the truncated
promoter compared to wt (Fig. 17H, lane 4 vs. lane 1). The requirement for HIF-1  in ET-1
transcription was maintained with the truncated ET-1 promoter reporter, as shown by PlGF
mediated increase in activity (Fig. 17H, lane 5 vs. lane 4) and by reduction of promoter activity
by both fenofibrate and exogenous miR-199a mimic (Fig. 17H, lanes 6 and 7). By contrast anti-
miR-199a augmented (~25%) the luciferase activity in fenofibrate treated cells, in the absence of
PlGF (Fig. 17H, lane 8). These data showed that KLF-11 did indeed participate in ET-1 gene
expression by repressing transcription under basal conditions and that ET-1 transcription is
subject to HIF-1  transactivation.

miR-199a2 levels were reduced in plasma of SCD vs. control human subjects
The status of miR-199a2 levels in human plasma samples was ascertained in an effort to
corroborate the in vitro results. As shown in Fig. 18A, levels of miR-199a-5p were significantly
lower by approximately 5-fold in plasma obtained from SCD subjects (SS, n=11) compared to
that obtained from unaffected sibling controls. As shown in Fig. 18B, the plasma ET-1 levels in
SCD samples were also higher compared to unaffected sibling control samples. We have
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previously shown that plasma ET-1 levels are significantly higher in SCD patients in comparison
to healthy controls (Sundaram et al., 2010; Yamashita et al., 2001). Thus the above results and
previously reported observations strongly indicate the existence of a reciprocal relationship
between miR-199a2 and ET-1, as was observed in vitro.

Effect of oral fenofibrate in wild type and Berkeley-sickle mice on miR-199a2 and ET-1
levels
Since the PPAR  agonist fenofibrate attenuated ET-1 expression in endothelial cells, we
examined the effect of the drug in vivo. BK-SS mice and C57BL6/6NJ wt control mice were fed
fenofibrate ad lib in the diet for 10 weeks. Lung tissues from these experimental animals were
analyzed for expression levels of premir-199a2, HIF-1α and ET-1 mRNAs, as described above.
A comparison of lung tissues from chow-fed diet vs. fenofibrate-chow diet to BK-SS mice,
showed a ~8-fold increase in miR-199a2 RNA (relative values 0.16 ± 0.08 vs. 1.25 ± 0.30), with
a corresponding decrease in HIF-1  mRNA levels and a significant reduction in ET-1 mRNA
levels (6.05 ± 0.22 to 0.07 ± 0.04) (Fig. 7C). In lung tissue obtained from wild type C57BL/6NJ
mice fed control chow diet vs. fenofibrate-chow diet, there was ~2.5-fold increase in miR-199a2
RNA expression (1.06 ± 0.27 to 2.58 ± 0.53), a decrease in HIF-1  mRNA and ~2-fold
reduction in ET-1 mRNA expression (1.03 ± 0.18 to 0.49 ± 0.19) (Fig. 18C). These data showed
fenofibrate fed to both BK-SS mice and wild type C57BL/6NJ mice augmented the expression of
miR-199a2 and was correlated with reduced ET-1 expression in lung, in support of our
regulatory model (Fig. 18G).
In order to further refine the lung gene expression data, we examined the levels of HIF-1 
mRNA and ET-1 mRNA in CD31
+
sorted endothelial cells from isolated lungs of control chow
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and fenofibrate-diet fed BK-SS mice. As shown in Fig. 18D, HIF-1  mRNA was significantly
reduced in isolated endothelial cells from fenofibrate-fed vs. control chow fed BK-SS mice.
Furthermore, ET-1 mRNA levels were also reduced in endothelial cells derived from fenofibrate-
fed, BK-SS mice vs. chow fed BK-SS mice (p=0.053) (Fig. 18E),
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Figure 18. miR-199a, HIF-1 , and ET-1 biomarkers in human SCD subjects and fenofibrate-treated
sickle mice. A. Plasma levels of miR-199a2-5p in SCD subjects (n=11) and healthy controls (n=13)
measured by specific TaqMan microRNA assay and normalized to RNU6B. B. Comparison of ET-1
levels in plasma of SCD subjects and healthy controls. C. Expression of premir-199a2, HIF-1α and ET-
1 mRNA in lung tissues obtained from C57B/6NJ and Berkeley SS mice fed either control chow diet or
fenofibrate chow for 10 weeks. RNA was extracted from lungs and qRT-PCR was performed on
samples (n=3-4) as indicated. D and E. Expression of HIF-1 mRNA and ET-1 mRNA in sorted
endothelial cells (CD31+) from lung tissue of fenofibrate-fed BK-SS mice and chow-fed BK-SS mice. F.
Kaplan-Meier survival plot of Berkeley SS mice (HbS; n=12) post fenofibrate feeding or control diet
feeding. G. Working model of miR-199a2 targeting 3’UTR of HIF-1α leading to reduction of HIF-1α and
concomitant reduction of ET-1 levels. miR-199a2 located within DNM3os is co-regulated by PPARα,
which binds to cis elements in DNM3os promoter. PPAR  agonist fenofibrate augments miR-199a2
expression leading to reduction in HIF-1α and ET-1.

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4.4 Discussion
We have previously shown plasma levels of placenta growth factor (PlGF), an angiogenic
growth factor, produced by erythroid cells, are high in SCD patients. This correlates with an
increase in plasma levels of ET-1 and tricuspid regurgitant jet velocity (TRV), the latter is a
marker of pulmonary hypertension (PH) in SCD (Sundaram et al., 2010). Similar to patients with
SCD, Berkeley sickle mice exhibit elevated plasma levels of PlGF and ET-1, and right
ventricular hypertrophy (Sundaram et al., 2010). Moreover, stimulated erythroid expression of
PlGF in normal mice, up to the levels seen in sickle mice, results in increased ET-1 production
and associated right ventricular hypertrophy with pulmonary changes within a time period of 8
weeks (Sundaram et al., 2010). These studies showed that PlGF-induced ET-1 expression in
vivo plays a significant role in development of PH in SCD.
Next, we examined the molecular mechanism of PlGF-induced expression of endothelin-
1 in cultured endothelial cells. Our previous studies show PlGF induces expression of ET-1 via
activation of hypoxia inducible factor-1  (HIF-1 ), independently of hypoxia. This involves
binding of HIF-1  to hypoxia response elements in the promoter of ET-1 (Patel et al., 2008).
In the present study, we examined the post-transcriptional mechanism of PlGF-mediated
ET-1 expression. We showed PlGF treatment of endothelial cells (HMEC) attenuated both
premir-199a2/premir-214 and mature miR-199a2/miR-214 levels. Based on several public data
sets, both miR-199a2 and miR-214 are predicted to target multiple genes. Previous studies show
miR-199a2 targets the 3’UTR of HIF-1  mRNA in mouse cardiac myocytes (Rane et al., 2009).
We focused our studies on the HIF-1  3’UTR as it harbors a complementary sequence to the
miR-199a2 seed sequence (positions +16 to +38 of the 3’UTR); HIF-1  mRNA has no
complementary sites for miR-214 binding. The specificity of miR-199a2 binding to the HIF-1 
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mRNA MRE was established using a luciferase reporter gene containing the HIF-1  3’UTR.
Synthetic miR-199a2 mimic attenuated basal and PlGF-induced luciferase reporter activity.
Moreover, the functionality of this sequence was established by mutation of the seed region
complement for miR-199a2 in the 3’UTR of the luciferase reporter. The mutated sequence
abrogated the inhibitory effect of synthetic miR-199a2 mimic and the stimulatory effect of anti-
miR199a2, thus confirming the presence of the miR-199a response element in the 3’UTR of
HIF-1  mRNA. These results also strongly implicated miR-199a2 involvement in post-
transcriptional regulation of HIF-1  expression in response to PlGF. Indeed, transfection of
HMEC with miR-199a mimic attenuated endogenous HIF-1  mRNA and protein, and
concomitantly attenuated basal and PlGF-induced ET-1 expression.
Previous studies have shown miR-199a is downregulated in cardiac myocytes in response
to low oxygen tension, which is required for the upregulation of target HIF-1  (Rane et al.,
2009). Hypoxia also regulates expression of several miRNAs that post-transcriptionally
modulate levels of a variety of mRNAs including HIF-1  (Nallamshetty, Chan, & Loscalzo,
2013). Indeed, our studies demonstrated that hypoxia attenuates miR-199a2 levels in endothelial
cells, which is manifested by increased HIF-1α expression as previously shown (Gonsalves &
Kalra, 2010).
miR-199a2 is located within the DNM3os transcription unit. This long non coding RNA
transcript is responsible for synthesis of the miRNA cluster containing miR-199a2 and miR-214,
and is involved in skeletal development (Desvignes, Contreras, & Postlethwait, 2014; Watanabe
et al., 2008). Thus it was necessary to determine whether PlGF regulated synthesis of the long
DNM3os transcript, as previously observed, or instead led to independent transcription of
premir-199a2. Our studies showed PlGF coordinately attenuated RNA levels of DNM3os,
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premir-199a2 and premir-214 in cultured human endothelial cells. As a consequence of reduced
miR-199a2 in PlGF-treated conditions, we expected increased levels of HIF-1  and ET-1
expression as was observed.
We observed significant reduction of miR-199a in plasma from patients with SCD compared
to unaffected sibling controls. Reduced levels of miR-199a in SCD would be expected to
decrease the turnover of HIF-1α mRNA leading to increased HIF-1  protein and subsequent
increased levels of ET-1, as was observed (Sundaram et al., 2010). Thus, both in vitro and in vivo
results support this model (Fig. 8E) whereby reduced expression of miR-199a2 leads to increased
expression of HIF-1α in SCD and is reflected by increased HIF-1  dependent gene expression.
Recent studies by others show myocardial hypoxia in mice, induced by transverse aortic
constriction, led to increased expression of DNM3os and concomitant miR-199a~miR214
expression (el Azzouzi et al., 2013). Herein, it was shown that miR-199a/miR-214 target PPARδ
mRNA for turnover, thus resulting in decreased PPARδ activity leading to defective
mitochondrial respiration, as a consequence of flawed fatty acid substrate metabolism. It was
suggested in their model, that miR-199a could also target HIF-1α mRNA for turnover. Reduction
of HIF-1  levels would lead to decreased DNM3os transcription and miRNA synthesis because
the promoter for the latter has HRE and Twist binding sites. Consequently the effects of hypoxia
are amplified because of this feedback loop (el Azzouzi et al., 2013). By contrast, our present
studies showed that hypoxia and PlGF in cultured endothelial cells reduced expression of
DNM3os, premir-199a2 and premir-214, resulting in higher expressed levels of HIF-1  and
increased expression of ET-1, the latter a key indicator of HIF-1  dependent transcription.
These results are consistent with previous studies which showed hypoxia upregulated ET-1
expression via activation of HIF-1  (Kourembanas, Marsden, McQuillan, & Faller, 1991;
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Yamashita et al., 2001). Further work is warranted to delineate the mechanism used during
hypoxia to regulate DNM3os transcription.
We examined the transcriptional regulation of miR-199a2 and its host gene DNM3os in
cultured endothelial cells. Utilizing PPAR  shRNA and PPAR  agonist, fenofibrate, we showed
PPAR  upregulated the expression of DNM3os, pre-mir199a2 and miR-199a2 RNAs. The effect
was specific for PPAR  since an antagonist to PPAR , GW6471, antagonized the stimulatory
effect of fenofibrate. In the promoter of DNM3os, there are three predicted PPAR  cis-binding
elements proximal to the transcription start site. These PPARα binding elements were involved
in DNM3os transcription as demonstrated by DNM3os-promoter-luciferase reporter assays and
response to PPARα agonist, fenofibrate. We refined our analysis of the PPARα cis-binding sites
in order to establish whether one or more of these PPARα elements were required for DNM3os
transcription in response to fenofibrate. Single PPARα site mutations were constructed for this
purpose whereupon it was found that two of the three PPAR  sites were required for PPARα-
dependent transcription of DNM3os, as determined by response to fenofibrate. These results
were further extended by analysis of physiological interactions of PPARα with the DNM3os
promoter in intact HMEC-1 as detected by ChIP assay. Thus the transcription activity of the
DNM3os locus under native conditions paralleled the transcription activity observed in reporter
assays.
The present study demonstrated for the first time to the best of our knowledge that
transcription factor PPARα co-regulated the transcription of DNM3os and miR-199a2. This
relationship between PPARα, HIF-1α and a downstream target gene, ET-1, are summarized in
our working model (Fig. 18G).
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Previous studies show that PPARα agonist, fenofibrate, attenuates ET-1 expression in a
PPARα dependent mechanism via activation of a Kruppel-like factor (KLF-11) (Glineur et al.,
2013) and by inhibition of AP-1 signaling pathways (Delerive et al., 1999). At this juncture, our
studies do not provide a link between PlGF-mediated activation of HIF-1α and modulation of
PPARα activity. Indeed there are contrary reports wherein HIF-1α has been shown to positively
regulate PPARα activity (Narravula & Colgan, 2001) in contrast to the opposing observation that
PPARα attenuates HIF-1α activity (Zhou et al., 2012). Tissue and cell specific functional
interactions between these two transcription factors may differ and have not been fully
addressed, thus further studies are warranted to resolve these opposing views. Nonetheless, we
observed PlGF mediated attenuation of DNM3os and pre-mir199a2 expression was linked to
ATF3, independently of PPARα. As we showed, PlGF increased ATF3 mRNA expression in a
time dependent manner, and transfection with ATF3 shRNA reversed PlGF-mediated attenuation
of DNM3os RNA levels, implying ATF3 acted directly as a repressor of DNM3os transcription.
Fibrates are FDA approved for the treatment of dyslipidemia (Lalloyer & Staels, 2010).
These PPAR  agonists have been shown to exhibit improvement of flow mediated dilation of
bronchial arteries in type 2 diabetes mellitus patients (Glineur et al., 2013) and diabetic
retinopathy in Type I diabetic animal models (Chen et al., 2013). Thus, we asked whether this
drug could be potentially beneficial in SCD treatment, given our experimental observation that
miR-199a2 synthesis was positively regulated by PPAR . As a first step, we examined the effect
of feeding fenofibrate to wild type control mice and Berkeley sickle mice. Results from these
studies demonstrated that 10 week administration of fenofibrate to these mice increased miR-
199a2 in lung tissue of both types of mice. There was a corresponding reduction of ET-1 mRNA
in lungs, consistent with the in vitro model, indicating that fenofibrate likely regulated the
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expression of ET-1 via miR-199a2 in vivo. Anecdotally, fenofibrate feeding also modestly
improved survival of Berkeley sickle mice (Fig. 18F).
In conclusion, our data showed transcription factor PPARα is capable of regulating the
expression of premir-199a2 and mature miR-199a2 via transcription of the host gene DNM3os.
In sickle subjects, miR-199a2 levels are significantly reduced, which in turn leads to increased
levels of HIF-1α and abnormal levels of ET-1; the latter of which is correlated as causal in PH
(Sundaram et al., 2010). These studies provide a mechanistic link for the transcriptional
regulation of miR-199a2 by fibrates, thus providing a therapeutic rationale for attenuating ET-1
levels in PH.

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CHAPTER 5: Placenta growth factor mediated transcription of
DNM3os/miR-199a2 is negatively regulated by ATF3 and associated histone
deacetylase 6 in sickle cell disease

5.1 Introduction
The onset of new transcription ascribed to PlGF is mediated mainly through HIF-1α.
Transcription of ET-1 and PAI-1 are among many genes of this regulon that are dependent on
HIF-1α for transcription, yet PlGF also participates in post-transcriptional regulation as well. We
have shown miRNAs directly post-transcriptionally regulate the expression of ET-1 and PAI-1
via targeting of the 3’-UTR of their mRNAs (Patel et al., 2011b) . Indirect, post-transcriptional
regulation of ET-1 and PAI-1 is achieved by miR-199a2, which targets the 3’-UTR of HIF-1α
mRNA (C. Li et al., 2014) (Chapter 4). Indeed the post-transcriptional regulation of HIF-1α
completes a regulatory loop that can modulate overall HIF-1α levels.
DNM3os produces a noncoding RNA that serves as the precursor of miR-199a2 and
miR-214 (C. Li et al., 2014) (Chapter 4). This locus is located within an intron of DNM3 and is
transcribed from the opposite strand, hence its designation. Transcription of DNM3os and co-
transcribed miR-199a2/miR-214 is attenuated by PlGF, thus allowing unhindered expansion of
HIF-1 α activity and expression of genes requiring this transcription factor, i.e. ET-1 and PAI-1
(C. Li et al., 2014) (Chapter 4). We found that positive regulation of DNM3os could be achieved
through peroxisome proliferator-activated receptor-α (PPARα) acting directly on the DNM3os
promoter(C. Li et al., 2014) (Chapter 4). These studies explained the molecular link between
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reduced miR-199a2 expression and increased expression of ET-1 in SCD subjects, as evident
from plasma levels of these biomarkers (C. Li et al., 2014) (Chapter 4). However, in SCD
subjects the molecular mechanism of PlGF mediated down-regulation of miR-199a2/miR-214
and its host gene DNM3os is not fully understood. Thus the basis for this dysregulation needs to
be clarified since this may be a major contributing factor in PH observed in SCD.
In the present study we show that repression of DNM3os in response to PlGF requires the
participation of activating transcription factor 3 (ATF3), which binds to ATF3 response elements
in the DNM3os promoter. ATF3, a stress-inducible gene, has been shown to play a role in
several pathological conditions such as host-defense immunity, cancer and hepatic
gluconeogenesis (Tsonwin Hai et al., 2010; J. Y. Kim et al., 2014; Wolford et al., 2013). To
delineate the mechanism by which ATF3 represses DNM3os transcription, we performed a
whole-cell proteomic analysis of ATF3 interacting proteins, identified by protein mass
spectroscopy of multi-protein complexes immunoprecipitated with antibody to ATF3. This
analysis revealed that ATF2, JDP-2, c-Jun, and HDAC6 co-immunopreicipated with ATF3 as
candidate factors responsible for repression of DNM3os.
Next, we explored whether chromatin remodeling was involved in transcriptional
repression. Histone deacetylation, mediated by histone deacetylases (HDACs), has been shown
to precede chromatin condensation, which prevents transcription factor access to DNA
regulatory elements and promoters thereby resulting in transcriptional repression (Cheung, Allis,
& Sassone-Corsi, 2000; Kouzarides, 2000). The participation of HDAC6 in regulation of
DNM3os was implicated by shRNA-mediated knockdownand Tubacin, a synthetic inhibitor of
HDAC6, both agents completely antagonized PlGF-mediated repression of DNM3os and miR-
199a2.
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5.2 Materials and Methods
Reagents
Recombinant human PlGF was purchased from Peprotech (Rocky Hill, NJ).; primary
antibodies to ATF3, JDP2 , ATF2 , c-Jun, HDAC6 and HDAC7, were obtained from Santa Cruz
Biotechnology (Santa Cruz, CA); anti-H3K9Ac and anti-H3K27Ac were obtained from Abcam
(Cambridge, MA); antibodies against β-actin and secondary antibodies conjugated to HRP were
purchased from Sigma Chemical Company (St. Louis, MO). The primers used for PCR
amplification of ATF3 promoter and its mutagenesis were purchased from Valuegene (San
Diego, CA). Unless otherwise specified, all other reagents were purchased from Sigma Chemical
Company (St. Louis, MO).
Mice
Berkeley sickle (BK-SS) mice originally obtained from Jackson Laboratories were bred
up to the 6th generation against a C57BL/6NJ background. BK-SS mice carry deletions for the
mouse α- and β-globin genes, and carry a transgene expressing the human β -globin and a mini
β-globin cluster including the β-sickle globin. In–house bred C57BL/6NJ animals served as
controls. Six to ten animals per group were used. Animal protocols were approved by the
Institutional Animal Care and Use Committee at Cincinnati Children’s Hospital Medical Center.
At the end of the study mice were exsanguiated and lungs were removed; tissues were stored at -
80
o
C for later assay of miRs and ET-1.

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RNA interference
The ATF3, JNK-1,JNK-2, PI3K, MAPK, HDAC 1,HDAC3, HDAC 4, HDAC 6, HDAC
7A, HDAC 8, HDAC 9, HDAC 11 and ATF2 shRNA clones were a generous gift from Dr. Jae
Jung (USC). The human shRNA library was purchased from Open Biosystems (now Thermo
Scientific, Grand Island, NY).The c-Jun and JDP2 siRNA was purchased from Santa Cruz
Biotechnology (Santa Cruz, CA); All synthetic shRNAs were supplied in a pGIPZ plasmid for
transfection.
Isolation of RNA and quantitative real-time-PCR (qRT-PCR) Isolation of RNA and
quantitative PCR
Total RNA was isolated from cells using TRizol reagent (Life Technologies, Grand
Island, NY) and mRNA expression was quantified using specific primers (Table 3) by
quantitative real time PCR (qRT-PCR) using the One-Step SYBR PrimeScript Kit (Clontech
Laboratories, Mountain View, CA) and the ABI Prism 7900HT sequence detection system
(Applied Biosystems, Foster City, CA)(C. Li et al., 2014) (Chapter 4).
Isolation and quantification of pre-miRNA and micro RNAs (miRNAs) by qPCR
microRNAs were purified utilizing the mirVana isolation kit (Life Technologies, Grand Island,
NY) according to the manufacturer's protocol. miRNA expression was quantified by qRT-PCR
and expression levels were normalized to a reference gene, RNU6B (U6 snRNA) (C. Li et al.,
2014) (Chapter 4). miRNAs were also quantified by Northern blot analysis utilizing biotinylated
probes for miR-199a2 and 5S rRNA, synthesized at Valugene (San Diego, CA) as described(C.
Li et al., 2014) (Chapter 4).
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Plasmids and plasmid construction
Luciferase Reporter vectors-- A 2 kb DNA segment containing the DNM3os promoter
was ligated into pGL3-basic luciferase reporter plasmid (C. Li et al., 2014) (Chapter 4)and used
Table 3. Oligonucleotides primers used in the Chapter 5 study.
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as the template for generating site-specific mutations. ATF3 site mutations were generated with
the Q5 Site-Directed mutagenesis kit (New England Biolabs, Ipswich, MA) utilizing primers
listed in Table 1. The resulting ATF3 mutant DNM3os promoters are designated ATF3mt1
(bases -216/-206), and ATF3mt2 (bases -789/ -779). Double mutations in the ATF3mt1 and
ATF3mt2 sites were generated from the ATF3mt1 construct as template using primers listed in
Table 1. All constructs and sequence mutations were verified by DNA sequencing (Retrogen,
San Di ego, CA). ATF3-luciferase promoter (bases -100/+35) and CRE mutated ATF3-luc
promoter (bases -100/+35) in pGL3 vector were kindly provided by Dr. Michael Kilberg
(University of Florida) (Fu & Kilberg, 2013)
Transient transfections
Endothelial cells (10
6
cells) were resuspended in 100 µ L of serum free RPMI-1640
medium containing indicated, appropriate shRNA vector and expression constructs (0.5μg), and
luciferase reporter plasmids (1.0μg), as indicated, followed by nucleofection utilizing appropriate
programs in the AMAXA nucleofector device (Lonza, Basel, Switzerland), as previously
described (J. Kang et al., 2009). The renilla luciferase plasmid (pRLSV40, 1.0 µ g) was co-
transfected with firefly luciferase reporter constructs to monitor transfection efficiency.
Following nucleofection, the cells were incubated in growth medium overnight, followed by
serum deprivation for 3 hrs, and treated with PlGF (250 ng/ml) for indicated time periods. The
cell lysates were assayed for luciferase activity using the Dual Luciferase Reagent kit (Promega,
Madison, WI). Luciferase values were normalized to renilla luciferase assay values and
expressed relative to the activity of the pGL3 control vector, as appropriate.

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Western blot analysis
Cells were lysed in RIPA buffer and 25 µ g of protein extracts were subjected to SDS-
PAGE, followed by Western blotting and chemiluminescence detection of bound antibody.
Membranes were probed with indicated dilutions of anti-ATF3(1:1000, sc-22798, Santa Cruz
Biotechnology, Santa Cruz, CA), anti-HDAC6(1:1000, sc-11420, Santa Cruz Biotechnology),
anti-HDAC7(1:500, sc-74563, Santa Cruz Biotechnology) and HRP- conjugated anti-rabbit
secondary antibodies (1:5,000, Sigma Aldrich, St. Louis, MO). Membranes were stripped and re-
probed with anti-β-actin (1:50,000, Sigma Aldrich). Quantitative analysis was performed using
the Image J analysis software.
Immunoprecipitation, Mass Spectroscopy and proteomic analysis
HMEC (1x10
6
cells) suspended in RPMI-1640 medium (0.1 ml) were transfected with
ATF3 expression plasmid (1 µ g) by nucleofection. The next day, cells were washed with PBS
and lysate prepared with RIPA buffer. Cell lysate from 2.5 x107 cells were incubated for 4 hr
with ATF3 antibody (Santa Cruz) in Tris-buffered saline (TBS) containing, 350mM NaCl and
0.3% Nonidet P40 (binding buffer), followed by the addition of Protein A/G agarose (Pierce)
and incubated overnight at 4° C on a rocking platform. The beads were washed six times with
binding buffer and antibody bound beads were subjected to reduction/alkylation (5 mM DTT at
56° C for 30 min then 25 mM Iodoacetamide in the dark 20 min), followed by treatment with 10
ng trypsin and lysC (modified sequencing grade, Roche) in sodium carbonate 50 mM buffer,
overnight at 37 ° C with shaking. After digestion the mixture was acidified with 10 μL of 10%
formic acid (FA) and peptides were recovered from the beads by filtering through C18 Stage Tip
column (Proxeon) followed by elution with 20 μL of 50% methanol/5% FA. The eluate was
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subjected to LC–MS/MS sequencing using an LC/MS system consisting of an Eksigent NanoLC
Ultra 2D (Dublin, CA) and Thermo Fisher Scientific LTQ Orbitrap XL (San Jose, CA). Protein
identifications were made using the commercially available Proteome Discoverer 1.4 (Thermo
Fisher Scientific) search engine.
Formaldehyde-assisted isolation of regulatory elements (FAIRE)
HMEC cells (5x10
6
cells) in complete media were treated with PlGF for 18 hrs followed
by the addition of formaldehyde to a final concentration of 1% and quenching with glycine as
described (Giresi, Kim, McDaniell, Iyer, & Lieb, 2007). Cells were scraped and subjected to
sonication for six, 18 s pulses, resulting in fragment lengths of ~300 to 400 bp; subsequent
procedural steps were followed as described in the protocol (Giresi et al., 2007). The genomic
(input) DNA and FAIRE DNA were isolated and purified, and subjected to quantitative PCR as
described in the protocol (Giresi et al., 2007) .
Chromatin immunoprecipitation (ChIP) assay
HMEC-1 (5x10
6
cells) cultured in 150 mm petri dishes were kept in SFM for 3 hr and
treated with PlGF (250 ng/ml) for 6 hr. ChIP analysis was performed using antibody to ATF3, c-
Jun, HDAC6, HDAC7, H3K9Ac, H3K27Ac and non-specific IgG (Santa Cruz Biotechnology,
Santa Cruz, CA) (Patel et al., 2009b). Briefly, immunoprecipitated DNA was air-dried and
resuspended in nuclease free water. DNA samples were subjected to PCR amplification utilizing
primers for the DNM3os promoter regions of interest corresponding to ATF3 binding sites and
distal site in ATF3 promoter (Table 1). PCR was performed for 35 cycles under the following
conditions: denaturation at 95° C for 30 s, annealing at 55° C for 60 s, and extension at 72° C for 2
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min. The PCR products were subjected to electrophoresis on a 2% agarose gel, visualized by
ethidium bromide staining, and quantified using the Image J analysis software.
Quantification of secreted ET-1
HMEC-1 (1x10
5
cells) were transfected with indicated shRNA in complete media and
incubated at 37° C for 24 hr. Cells were washed with serum-free medium (SFM) and incubated
for 3 hr in SFM (2 ml). Treatments were begun by replacement of SFM with fresh SFM (1 ml).
Cells were treated with either PlGF (250 ng/ml), shRNAs or Tubacin) overnight. The culture
supernatant was collected and an aliquot (0.1 ml) was assayed for ET-1 using an enzyme-linked
immunosorbent assay (ELISA) kit (Assay Design/Enzo Life Sciences, Farmingdale, NY) (Patel
et al., 2008). The cells were isolated by scraping and the pellet was assayed for protein content
utilizing the Bradford method. Plasma specimen ET-1 levels were determined using DuoSet
ELISA kit from R&D (Minneapolis, MN)
Statistical analysis
The significance between two data groups was ascertained using an unpaired Student’s t-
test, and results are presented as means ± S.E.M. * P<0.05, ** P<0.01, *** P <0.001, and ns (not
significant) with P>0.05

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5.3 Results
PlGF up regulates expression of ATF3, which involves activation of PI-3-kinase, MAP
kinase and Jun-N-terminal kinase -1 (JNK-1).
In the previous chapter we showed that PlGF treatment of cultured human endothelial
cells repressed DNM3os and miR-199a2 transcription (Chapter 4). However, this mechanism
remains elusive. As shown in Fig. 19A, PlGF treatment of HMEC-1 resulted in a time dependent
increase in ATF3 mRNA expression, with the maximum increase of ~3.5-fold observed at 2 hr.
At 2 hr there was also ~1.5-fold increase in ATF3 protein expression (Fig.19B). Transcription of
DNM3os during PlGF treatment declined by 50% at 6 hr.
Next, we examined the intracellular signaling pathway for induction of ATF3, utilizing
pharmacological inhibitors specific for signal transduction protein kinases. We utilized the
established kinase inhibitors LY 294002 (PI-3 kinase), PD98059 (MAP kinase), SB203580
(p38MAP kinase) and SP600125 (JNK). As shown in Fig. 18C, these drugs completely inhibited
PlGF-induction of ATF3 mRNA and protein expression to at or below basal levels, Fig. 18C and
Fig.18D, respectively.

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Figure 19. PlGF mediated expression of ATF3 involves signaling via PI-3-Kinase, MAP kinase,
p38MAPK and Jun-terminal kinase-1. A, Time course of PlGF-mediated expression of ATF3 and c-Jun
mRNA. B, PlGF-mediated expression of ATF3 protein at 2hr. C and D, Effect of pharmacological
inhibitors for specific kinases on PlGF-mediated ATF3 mRNA and protein expression, respectively. E,
Effect of shRNAs for PI-3 kinase, MAPK, JNK-1 and JNK-2 on PlGF-mediated ATF3 mRNA expression.
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In order to minimize the possibility of non-specific inhibitor effects, we utilized
exogenous dominant negative Dn-PI-3kinase, MAPK shRNA, JNK-1 shRNA and JNK-2 shRNA
to inhibit the respective protein kinases in HMEC. As shown in Fig. 18D, we observed that Dn-
PI-3-kinase and shRNAs for MAPK and JNK-1 completely abrogated PlGF induction of ATF3.
The inhibitory activity of SP600125 affects all JNKs, thus it was necessary to further resolve the
effect of this inhibitor. Silencing of JNK-1 and JNK-2 was achieved using specific exogenous
shRNAs. As shown in Fig. 18E, JNK-2 shRNA had no effect on PlGF induction whereas JNK-1
shRNA treatment allowed only a two-fold increase in ATF-3 mRNA in response to PlGF
induction. Taken together, these data showed PlGF mediated induction of ATF3 mRNA
involved PI-3Kinase, p38MAP kinase, MAP kinase and JNK2.

PlGF-mediated upregulation of ATF3 results in transcriptional repression of pre-miR-
199a2 and its host gene DNM3os
Next, we examined whether ATF3 was involved in repression of DNM3os transcription.
As it was shown previously in Chapter 4, the DNM3os transcription unit includes miR-199a2
and miR-214, implying that transcription of DNM3os leads to synthesis of a pri-miR-199a2/214
precursor that is further processed into discrete miRNA molecules. We utilized a loss and gain of
function approach to analyze the role of ATF3 in regulation of DNM3os transcription. Under
basal conditions, in the absence of PlGF treatment, transfection with ATF3 shRNA had no effect
DNM3os transcription, similar to that observed with scrambled (sc) RNA, (Fig. 19A, lane 2 vs.
lane 3). Synthesis of pre-miR199a2 also was unaffected by ATF3 shRNA (Fig. 19B, lanes 2 vs.
lane 3); however, upon induction with PlGF there was ~70% reduction in RNA synthesis from
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DNM3os (Fig. 19A, lane 4 vs. lane 1) and a corresponding ~60% reduction of pre-miR199a2
(Fig. 19B, lane 4 vs. lane 1). The repression observed by PlGF treatment was reversed upon
transfection with ATF3 shRNA, compared to scRNA, as indicated by increased DNM3os RNA
synthesis (Fig. 19A, lane 5 vs. lane 6) and expression of pre-miR199a2 (Fig. 19B, lane 5 vs. lane
6). These data showed ATF3 participated in repression of DNM3os transcription with
concomitant effect on miR-199a2 synthesis.
As further validation of ATF3 acting as a repressor of DNM3os transcription, HMEC-1
were transfected with wt ATF3 expression plasmid (ATF3 variant 1) and a truncated ATF3
expression plasmid (DN-ATF3), lacking the DNA binding domain. In the absence of PlGF,
expression of wt ATF3 reduced basal levels of DNM3os (Fig. 19C) and pre-miR199a2 (Fig.
19D) by ~70%, whereas DN-ATF3 was ineffective under these conditions (Fig. 19C and 19D).
These results showed that functional ATF3 was required for repression of DNM3os
transcription.
Repression of DNM3os transcription in HMEC-1 was also confirmed by measuring miR-
199a2 synthesis by Northern blotting following various treatments. As shown in Fig. 19E, PlGF
treatment resulted in ~70% reduction of mature miR-199a (Fig. 19E, lane 2 vs. lane 1), which
was consistent with the observed qRT-PCR results (Fig. 19A). Pre-incubation of these cells with
ATF3 shRNA prior to PlGF induction partially antagonized the expected repressive effect, where
only a 20% reduction of miR-199a2 level was observed. Transfection of wt ATF3 plasmid, in the
absence of PlGF, mimicked the repression observed with the latter and resulted in a 60%
reduction of miR-199a2. This product analysis confirmed that repression of the DNM3os locus
(including miR-199a2) following PlGF treatment also involved ATF3, possibly as part of a
repressor complex.
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Figure 20. ATF3 acts as repressor of DNM3os and miR-199a2 transcription. A and B, Effect of ATF3
shRNA on PlGF- mediated DNM3os and pre-mir-199a2 RNA expression. C and D, Effect of ATF3
expression via ATF3 expression plasmid (variant 1) or truncated ATF3 plasmid on DNM3os and pre-
miR-199a2 RNA expression. E, Northern blot of miR-199a2 expression in HMEC in response to
treatment with PlGF, transfection with ATF3 shRNA or overexpression of ATF3 variant 1.
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ATF3 acts as a repressor of DNM3os as demonstrated by DNM3os promoter analysis and
chromatin immunoprecipitation assay
In silico analysis of the DNM3os 5’-flanking region (~2.0 Kb) revealed the presence of
seven potential ATF3 cis-binding elements, the two sites proximal to the transcription start site
are depicted in the promoter schematic of Fig. 20A. In order to firmly establish the role of ATF3
in the regulation of DNM3os, we employed a wt DNM3os promoter-luciferase construct as a
model for further analysis. Transfection of HMEC-1 with wt DNM3os-luciferase reporter
plasmid followed by PlGF treatment, reduced luciferase activity by ~60% (Fig. 20B, lane 2 vs.
lane 1). Co-transfection of the reporter with ATF3 shRNA reversed the reduction of luciferase
activity (Fig. 20B, lane 3 vs. lane 2), while the effect of scRNA was not significant (Fig. 20B,
lane 4 vs. lane 2). This indicated the reporter construct behaved similarly to the endogenous
DNM3os promoter with respect to PlGF mediated repression and was judged to be valid for
further analysis.
Next, we generated base substitution mutations in the wt DNM3os reporter luciferase
plasmid, wherein ATF3 -binding sites proximal to the transcription start site of the DNM3os
promoter were mutated as depicted in schematic of Fig. 20A. In HMEC-1 transfected with
reporters with single mutations at site 1 (ATF3 mt1, Fig. 20B, lanes 5 and 6 vs. lane 4) or site 2
(ATF3 mt2, Fig. 20B, lanes 9 and 10 vs. lane 8), neither PlGF treatment nor co-transfection with
ATF3 shRNA affected luciferase activity. Moreover, transfection of HMEC-1 with the double
ATF3 site mutations of the DNM3os promoter (ATF3 DM) exhibited no change in luciferase
activity in response to either PlGF or ATF3 shRNA (Fig. 20B, lanes 12 and 13 vs. lane 11).
Taken together, these results showed ATF3 binding to the DNM3os promoter participated in
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PlGF-dependent repression of DNM3os transcription and both ATF3 binding site 1 and site 2 are
important for the recruitmet of regulatory elements.
Next, we examined the effect of exogenous ATF3 on DNM3os reporter activity. As
shown in Fig. 20C, inclusion of the ATF3 expression plasmid, in the absence of PlGF, reduced
wt DNM3os reporter activity by ~60% while scRNA had no significant effect (Fig. 20C). Taken
together, these data showed ATF3 acted as a repressor for DNM3os transcriptional activity.
The activity studies regarding the effect of ATF3 on DNM3os promoter activity were
molecularly characterized by chromatin immunoprecipitation analysis (ChIP). Occupancy of
ATF3 on the endogenous DNM3os promoter was also compared between cells transfected with
ATF3 shRNA and cells supplemented with exogenous ATF3. Chromatin samples were
immunoprecipitated with ATF3 antibody and analyzed for DNA recovery by RT-PCR. HMEC-1
transfected with ATF3 expression plasmid showed ~2-fold increase in the expected PCR product
size for site 1 (91 bp), corresponding to the DNM3os promoter region, TSS proximal site (nt -98
to -8; Fig. 20D (left), middle lane vs. lane 1). Conversely, transfection with ATF3 shRNA
reduced ATF3 binding to site 1 (Fig. 20D (left), lane 3 vs. lane 1). Similarly, chromatin obtained
from HMEC-1 transfected with ATF3 shRNA showed reduced binding of ATF3 to site 2 in the
DNM3os promoter, as shown by reduced recovery of the expected PCR product (115 bp; Fig.
20E), corresponding to nt -912 to -798 of the DNM3os promoter, containing the second ATF3
site. The amplification of input DNA before immunoprecipitation of chromatin samples (Figs.
20D and E, middle rows) was equivalent in all samples. Immunoprecipitation of chromatin
samples with control rabbit IgG did not recover any DNA with the expected PCR product sizes
(Figs. 20D and E, bottom rows). These data showed ATF3 binding to the DNM3os promoter was
functionally correlated with conditions promoting repression of DNM3os transcription.
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117

Characterization of accessory proteins of ATF3 involved in transcriptional repression of
DNM3os by Proteome Analysis
Previous work has established that ATF3 can act as either a repressor or activator of
transcription; this functional difference is mediated by recruitment of partner proteins, e.g. c-Jun
dimerization protein (JDP2), ATF2, AP-1 complex and histone deacetylases (HDACs) (Darlyuk-
Saadon et al., 2012). In order to understand the functional role of ATF3 in repression of
DNM3os, we examined its recruitment of additional protein(s) in response to PlGF.
HMEC-1 cells were exogenously enriched in ATF3 by transfection of an ATF3
expression plasmid. This approach mimicked PlGF-induced ATF3 expression as was previously
observed (Fig. 18A) and was found to similarly repress DNM3os under basal conditions.
HMEC-1 cells were transfected with ATF3 expression plasmid and incubated overnight prior to
cell lysate preparation and immunoprecipitation with ATF3 antibody. The purified
Figure 21. ATF3 transcription factor binding to cis-binding elements in the promoter of DNM3os. A,
Schematic representation of ATF3 cis-binding elements in the promoter of DNM3os, and exhibiting
mutations in core sequences corresponding to ATF3 sites (1 and 2). B. Effect of PlGF and ATF3shRNA
on wt. DNM3os, ATF3 mutant1 and ATF3 mutant 2 reporter luciferase activity. HMEC were transfected
with either DNM3os (wt) –luc plasmid or the mutant plasmid with singly mutated ATF3 binding sites (1
or 2) in DNM3os-luc constructs (ATF3mt); Transfected cells were treated as indicated with PlGF for 24
hr. The cell lysates were assayed for luciferase activity and normalized to Renilla activity. C. Effect of
expression of ATF3 via ATF3 variant 1 expression plasmid on wt DNM3os reporter luciferase activity.
D. ChIP analysis of ATF3 binding to native chromatin. HMEC cells were transfected overnight with
either ATF3shRNA or ATF3 variant1 expression plasmid. The soluble chromatin was isolated and
immunoprecipitated with either antibody to ATF3 or control rabbit IgG. The primers employed flank
ATF3 binding sites (1 and 2), as indicated in schematic shown in Fig. 20A, and are listed in Table 1.
Immunoprecipitated chromatin was processed for PCR analysis and expected product sizes are
indicated. The bottom panel shows amplification of the input DNA before immunoprecipitation.
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immunocomplexes were subjected to proteolytic digestion followed by mass spectroscopy and
proteomic analysis.
This analysis identified ATF3 immunocomplexes were formed with ~84
proteins/polypeptides; the highest scoring candidates are ATF2, ATF3, JDP2, HDAC6, HDAC7,
JNK1, and c-Jun, as listed in Table 4. Based on a gene ontology bias, we focused our analysis of
proteins which may be involved in transcriptional repression or activation.

Table 4. Proteomic analysis of HMEC cells transfected with ATF3 expression plasmid, followed by
immunoprecipitation with ATF3 antibody and mass spectroscopic analysis
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Histone deacetylase (HDAC)
participates in PlGF mediated
repression of DNM3os RNA
transcription
Histone deacetylases (HDACs) are
associated with repression of gene
transcription; these act by histone
deacetylation and thereby promote
chromatin condensation (Narlikar, Fan, &
Kingston). The HDAC family comprises
four enzyme classes: class 1 (HDAC 1, 2,
3 and 8), class II (class IIa: HDAC5, 7 and
9); class IIb: HDAC6 and 10), class III
(sirtuin 1-7) and class IV (HDAC11)
(Narlikar et al.). We examined possible
roles of HDAC 1, 3, 4, 6, 7, 8, 9 and 11 in
PlGF-mediated repression of DNM3os,
utilizing available HDAC shRNAs. As
shown in Fig. 21A, shRNAs for HDACs 6
and 7, but not the other HDACs tested,
antagonized PlGF-mediated repression
of DNM3os. Since pre-miR-199a2 is
part of the DNM3os transcription unit,
Figure 22. Association of histone deacetylase in
chromatin remodeling and transcription of DNM3os/miR-
199a2. A and B, Effect of shRNAs for HDACs (1, 3, 4, 6,
7, 8, 9 and 11) on DNM3os and pre-miR-199a2 RNA
levels. C. Effect of two different clones of HDAC6 shRNAs
on corresponding ptrotein expression.
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we examined the synthesis of pre-miR199a2 mRNA following transfection with HDAC
shRNAs. As expected, the expression of pre-miR-199a2 was decreased by HDAC 6 and 7
shRNAs, but not by other HDAC shRNAs (Fig. 21B). The activities of HDAC 6 shRNAs were
confirmed to knockdown expression of the respective proteins as shown by western blots; the
shRNAs were ~80-90% effective in attenuating corresponding protein synthesis (Fig. 21C).
Since PlGF treatment did not change HDAC6 mRNA levels, through 6 hr following PlGF
treatment, another mechanism is likely responsible for increased nuclear activity or localization
of these enzymes with DNM3os. These data clearly showed HDAC6 were involved in PlGF-
mediated repression of DNM3os RNA and pre-miR-199a2.

Tubacin, a selective inhibitor of HDAC6, reverses PlGF mediated repression of DNM3os
transcription
In order to further characterize HDAC participation in DNM3os repression we utilized
chemical inhibitors selective for HDACs for their effects on basal and PlGF mediated
transcription of DNM3os. Currently, selective chemical inhibitors are available for only a few
HDACs. We tested the effect of trichostatin (TSA), a broad spectrum HDAC class I and II
inhibitor, Tubacin, a specific inhibitor of HDAC6, and Mocetinostat (MGCD0103) a selective
inhibitor for HDAC1, with no inhibitory activity for HDAC 4, 5, 6, 7 and 8. A dose dependent
effect of Tubacin on derepression of DNM3os transcription was observed with optimal effect at
5 µ M (Fig. 22A). A comparison of the effects Tubacin on DNM3os transcription with the other
deacetylase inhibitors was performed. Both TSA and Tubacin completely reversed PlGF-
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mediated repression of DNM3os transcription (Fig. 22B, lanes 3 and 4 vs. lane 2), while
Mocetinostat or vehicle (DMSO) control had no effect (Fig. 22B, lanes 5 and 6 vs. lane 2).

Figure 23. Effect of histone deacetylase inhibitors on DNM3os transcriptional activity. A, Dose response
effect of Tubacin, an HDAC6 inhibitor, on DNM3os RNA expression. B, Effect of Trichostatin (TSA-
HDAC pan inhibitor), Tubacin (HDAC6 inhibitor) and Mocetinostat (HDAC pan inhibitor) on DNM3os
RNA expression. Cells were incubated with inhibitors for 6hr. C. Effect of HDAC inhibitors on DNM3os
transcription activity as measured by reporter luciferase activity.
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Is ATF3 binding to the DNM3os promoter essential for repression?
Since knockdown of both ATF3 and HDAC6 by shRNAs reversed PlGF mediated
repression of DNM3os, we examined this further utilizing a luciferase reporter driven by the
DNM3os promoter. In particular we were interested in the binding of either ATF3 or HDAC6 to
this promoter as a co-dependent or mutually exclusive process for repression. As shown earlier
(Fig. 20B) expression from this reporter was repressed by PlGF like the endogenous promoter.
This indicated to us that the reporter plasmid was chromatinized and would respond to chromatin
remodeling activities.
PlGF treatment of HMEC-1 transfected with the DNM3os promoter reporter resulted in
40% inhibition of luciferase expression. The repression was reversed in the presence of either
TSA or Tubacin, which was consistent with the effect of these inhibitors on the endogenous
DNM3os promoter (Fig. 22C).
The importance of the ATF3 binding sites in the DNM3os promoter were examined as
necessary elements needed for DNM3os repression. HMEC-1 cells were transfected with the
DNM3os luciferase reporter containing mutations of both distal and proximal ATF3 binding sites
relative to the TSS. As shown in Fig. 22C, loss of both ATF3 binding sites of the DNM3os
luciferase reporter rendered the promoter insensitive to PlGF mediated repression and
furthermore both TSA and Tubacin potentiated the effect of PlGF. (Fig. 22C) These results
indicated that ATF3 sites in the DNM3os promoter were required for PlGF mediated repression
of DNM3os transcription. The effects of the histone deacetylase inhibitors implicated HDAC6 as
a participant in the repression of DNM3os, likely by a PlGF mediated signal for chromatin
remodeling and repression of DNM3os transcription.
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Binding of ATF3, c-Jun and HDAC6 to the DNM3os promoter as demonstrated by
chromatin immunoprecipitation (ChIP)
In order to gain information regarding DNA binding proteins present on the DNM3os
promoter in response to PlGF mediated repression, we examined whether transcription factors
ATF3, JDP2, ATF2, c-Jun and HDACs were present on the promoter at locations centered near
the two identified ATF3 binding sites. For this analysis we performed chromatin
immunoprecipitation analysis of chromatin isolated from PlGF treated cells.
As shown in Fig. 23A, we observed significant binding of ATF3 and HDAC6 to the
DNM3os promoter in response to PlGF. Both ATF3 and HDAC6 showed ~2-fold increased
binding centered on ATF3 site 1 with ~50% increased binding at ATF3 site 2, while JDP2
showed a ~2-fold increased binding on ATF site 1 and a ~4-fold increased binding at ATF3 site
2. DNA recovery with anti-HDAC7 showed a modest 20% increase at both ATF3 sites. Binding
of c-Jun and ATF2 was not significantly changed in response to PlGF at either ATF3 site (Fig.
23A, Panel 1 and 2).
As a control, a distal region from the DNM3os gene was also analyzed by ChIP with the
same set of antibodies. The resulting PCR analysis revealed no DNA recovery from this distal
site by any of the factor specific antibodies (Fig. 23A Panel 3). Moreover, there was equal input
of DNA in both untreated and PlGF treated samples as shown in Fig. 23A (Panel 1 and 2).
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Taken together, the data showed PlGF mediated repression of DNM3os transcription
likely involved association of ATF3, HDAC6 and JDP2 on the endogenous promoter centered
near ATF3 sites.

Figure 24. Binding of transcription factors, ATF3, JDP2, c-Jun, HDAC6 ATF2 and HDAC7 to DNM3os
promoter in native chromatin as assessed by ChIP. HMEC cells were treated with PlGF for 2 h. The
soluble chromatin was isolated and immunoprecipitated with test antibody or control rabbit IgG. The
PCR primers flanking ATF3 binding sites (1 and 2) in DNM3os promoter, as indicated in schematic
shown in Fig. 20A, and listed in Table 1. Immunoprecipitated chromatin was processed for PCR
analysis and expected product sizes are indicated. The bottom panel shows amplification of the input
DNA before immunoprecipitation. Panel 3, Immunoprecipitated chromatin was processed for PCR
analysis utilizing primers corresponding to non ATF3 binding site in the DNM3os promoter.
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Accessibility of transcription factors in gaining access to chromatin and regulating
DNM3os gene expression (FAIRE)
In order to better understand the consequence of trans-acting factor binding to the
DNM3os promoter during PlGF mediated repression, we examined the chromatin status of this
gene. For this analysis, we utilized FAIRE (formaldehyde assisted isolation of regulatory
elements) methodology, which identifies nucleosome-free, euchromatic regions of the genome
(Giresi et al., 2007), although in the present study we examined the DNM3os locus. HMEC-1
were used for FAIRE-qPCR analysis following treatment in the presence or absence of PlGF
induction. Cells treated with PlGF showed a ~80% reduction of FAIRE signal at ATF3 sites 1
and a ~70% reduction at ATF3 site 2, compared to untreated cells (Fig. 24A and B, lane 2 vs.
lane 1); this was likely due to chromatin remodeling, consistent with the observed repression of
transcription.
Conversely, treatment with Tubacin in the presence of PlGF increased the FAIRE signal
~9-fold at ATF3 site 1 and ~5-fold at ATF3 site 2, due to opening of the chromatin structure
(Fig. 24A and B, lane 3 vs. lane 2), compared to the PlGF treatment condition. HMEC
transfected with HDAC6 shRNA showed ~6-fold higher FAIRE signal at ATF3 site 1 and ~3-
fold higher FAIRE signal at ATF3 site 2 compared to PlGF treatment (Fig. 24A, lane 5 vs. lane
2). Furthermore, the FAIRE signal was ~3-fold higher with shRNA for ATF3 at ATF3 site 1 and
~6-fold higher at ATF3 site 2 compared to PlGF treatment alone, indicating that shRNA for
ATF3 helped retain open chromatin structure in the presence of repressive PlGF (Fig. 21A lane
4 vs. lane 2). Taken together, the data showed PlGF mediated upregulation of ATF3 causee
chromatin condensation at cis-regulatory regions in the DNM3os promoter to repress DNM3os
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transcription. Conversely, ATF3 shRNA and HDAC6 shRNA induced chromatin opening at or
adjacent to cis-regulatory regions in the DNM3os promoter to increase DNM3os transcription.

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De-acetylation of histone H3 marks in the promoter of DNM3os is required for repression
In silico analysis using ENCODE data of the DNM3os gene, located on chromosome 1,
showed extensive histone acetylation marks for H3K27Ac in a ~1.2 kb region centered on the
TSS. Thus, we examined the presence of H3K9Ac and H3K27Ac marks in the promoter of
DNM3os proximal to the identified ATF3 sites under basal conditions and PlGF mediated
repression. These histone marks are associated with chromatin remodeling events occurring at
promoter and enhancer binding (Jenuwein & Allis, 2001). Chromatin was isolated from HMEC-
1 treated with PlGF followed by immunoprecipitation with either anti-H3K9Ac or anti H3K27Ac
antibodies. The recovered DNA was analyzed by PCR using primers corresponding to the
respective ATF3 binding sites (see primer listing in Table 1). As shown in Fig. 24C, PlGF
reduced by ~50% the recovery of DNA with anti H3K9Ac proximal to ATF3 site1. DNA
recovery was enhanced by inclusion of Tubacin, HDAC6 shRNA or ATF3 shRNA. Consistent
with this result, PlGF treatment reduced by ~75% DNA recovery proximal to ATF3 site 2.
Similar to ATF3 site 1, DNA recovery was enhanced by Tubacin, HDAC6 shRNA or ATF3
shRNA to levels observed in untreated cells (Fig. 24D). For comparison DNA recovery with
anti-H3K27Ac was also examined, as this modification is also involved in chromatin
remodeling. As shown in Fig. 24C, PlGF treatment reduced recovery of DNA by ~65%,
proximal to ATF3 site 1 of the DNM3os promoter; DNA recovery was enhanced by treatment
Figure 25. Effect of Tubacin, ATF3 shRNA and HDAC6 shRNA on chromatin structure of ATF3
binding site 1 and 2. A and B. FAIRE-qPCR on ATF3 site 1 and site 2 in the presence of PlGF, in
Tubacin treated, ATF3 shRNA and HDAC6 shRNA transfected HMEC-1. C and D. ChIP-qPCR with
H3K9Ac, H3K27Ac and IgG antibody on ATF3 site 1 and site 2 in the presence of PlGF, in Tubacin
treated, ATF3 shRNA and HDAC6 shRNA transfected HMEC-1.
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with Tubacin, HDAC6 shRNA and ATF3 shRNA. Similarly, PlGF attenuated DNA recovery by
~80% proximal to ATF3 site 2 of the DNM3os promoter. DNA recovery was increased by ~50%
by co-addition of Tubacin, HDAC6 shRNA, and ATF3 shRNA (Fig. 24D). Control incubations
with non-specific, primary rabbit IgG were negative for DNA recovery and chromatin
immunoprecipitation from a TSS distal site with anti-H3K9Ac or anti-H3K27Ac antibodies
showed no changes upon PlGF addition.
These results were interpreted to indicate that chromatin remodeling of the DNM3os
promoter region occurred during the transition from basal expression to the repressed state
following PlGF induction. The presence of H3K27Ac and H3K9Ac histone marks are
characteristic of active chromatin, thus the effects of Tubacin and HDAC6 shRNA effectively
antagonized this change in chromatin structure in the DNM3os promoter encompassing the two
ATF3 binding sites. The role of ATF3 is less clear since as it may be required for guiding
chromatin remodeling enzymes (e.g. HDAC6) and/or other accessory proteins to effect the
silencing of this promoter.

HIF-1α -dependent transcription can be modulated by ATF3 levels: effects on ET-1
expression
Previous studies show PlGF induces HIF-1α expression, independently of hypoxia, which
in turn augments ET-1 expression (Patel et al., 2008), thus we examined whether ATF3 activity
was correlated with ET-1 mRNA and protein expression. HMEC-1 cells were transfected with
ATF3 shRNA followed by treatment with PlGF. Under these conditions, PlGF-induced HIF-1α
and ET-1 mRNA expression were antagonized by ATF3 shRNA (Fig. 25A, lane 3 vs. lane 2).
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Next, we examined whether ATF3 affected release of ET-1 protein from HMEC-1 cells. As
shown in Fig. 25B, PlGF treatment of HMEC-1 showed ~1.8 fold increase in the secretion of
ET-1 protein (Fig. 25B, lane 2 vs. lane1). Transfection of HMEC-1 with ATF3 shRNA followed
by PlGF treatment reduced ET-1 secretion to the basal level (Fig. 25B, lane 3 vs. lane 2).
Conversely, expression of ATF3 plasmid, but not ATF3 truncated plasmid lacking the DNA
binding domain (DN-ATF3), in the absence of PlGF, led to increased ET-1 secretion (Fig. 25B,
lane 4 vs. lane 5). Taken together, the data showed attenuation of ATF3 levels could be
correlated with decreased ET-1 secretion, while increased expression of ATF3 was correlated
with increased secretion of ET-1 protein from HMEC cells. Based on the proposed regulatory
model, ATF3 levels are transduced through changes in DNM3os transcription and miR-199a2
synthesis, thus modulating HIF-1α mRNA and subsequent target gene transcription.

Chromatin status indirectly affects ET-1 expression and secretion
The DNM3os locus was demonstrated to undergo repression by a change in chromatin
state/status. As shown above, HDAC6 activity was important for ATF3 mediated repression of
DNM3os/miR-199a2 expression. Consequently, we asked whether Tubacin affected expression
of ET-1 by this process. As shown in Fig. 25C, PlGF induced secretion of ET-1 was attenuated
by derepression of DNM3os transcription resulting from Tubacin treatment or by inclusion of
HDAC6 shRNA. Thus the effects of these treatments were consistent with the modulation of
HIF-1α levels mediated by changes in miR-199a2.
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Figure 26. A. qRT-PCR of HIF-1α and ET-1 mRNA in PlGF treated HMEC and with transfection of
ATF3 shRNA. B and C. ET-1 section measured by ELISA, in the presence of PlGF. D. lung ATF3
RNA level in BKSS and C57 mice. E. Western blotting of ATF3 protein in the mice lung tissue and
quantification of ATF3/β-actin is shown in panel F.
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ATF3 expression in lung tissues of Berkley Sickle Cell mice (BK-SS)
Since PlGF mediated upregulation of ATF3 leads to repression of miR-199a2 and
concomitant upregulation of ET-1 expression in vitro, we examined the expression of ATF3 in
BK-SS mice. These animals exhibit abnormally high PlGF levels compared to control
C57BL/6NJ mice. As shown in Fig. 25D, measurements of ATF3 mRNA levels in lung tissues,
the site of PHT in BK-SS mice, showed higher levels compared to control C57BL/6NJ mice.
Moreover, ATF3 protein levels were also higher in lung tissues of BK-SS mice compared to
control C57BL/6NJ mice (Fig. 25E and F). As previously reported, in comparison to control
animals, lung tissues of BK-SS mice show lower levels of pre-miR-199a2 with higher levels of
ET-1 mRNA (C. Li et al., 2014). Taken together, these data nicely correlate higher PlGF levels
with increased expression of ATF3, both in vitro and in vivo. Thus changes in ATF3 levels can
be manifested by changes in miR-199a2 levels with subsequent effect on HIF-1α and associated
gene regulation. The data obtained from BK-SS mice indicated that this mode of regulation is
likely dysfunctional in SCD.
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5.4 Discussion
Previous studies in cultured endothelial cells showed PlGF induces expression of ET-1
via activation of HIF-1α, independently of hypoxia (Patel et al., 2008). This involves binding of
HIF-1α to hypoxia response elements in the promoter of ET-1. Moreover, our studies show PlGF
attenuates RNA levels of pre-miR-199a2 and concomitantly mature miR-199a2 in vitro (Chapter
4). In Chapter 4, we showed that in BK-SS sickle mouse model and in SCD patients, the plasma
levels of miR-199a2 are significantly reduced compared to normal controls. This observation is
significant because our recent studies show miR-199a2 targets the 3’-UTR of HIF-1α mRNA and
attenuates HIF-1α expression (Chapter 4).
In light of this regulatory loop, a consequence of PlGF-treatment of HMEC-1 cells and
abnormally high PlGF levels as seen in the SCD mouse model, we anticipated increased levels of
HIF-1α with ET-1 expression, as was observed (Chapter 4). Both in vitro and in vivo results
showed a consequence of reduced expression of miR-199a2 is increased expression of HIF-1α,
especially in SCD, as reflected by increased HIF-1α dependent gene expression.
Transcription of DNM3os produces a long noncoding RNA which is the primary
transcript for miR-199a2 and miR-214 (el Azzouzi et al., 2013; C. Li et al., 2014; Loebel et al.,
2005). Although this locus is positively regulated by Twist during embryogenesis (Loebel et al.,
2005), we demonstrated that it is subject to additional positive regulation in SCD. We have
extended our previous studies by demonstrating that DNM3os transcription is also subject to
negative regulation mediated by PlGF. We provided evidence that PlGF-mediated induction of
activating transcription factor 3 (ATF3) expression participated in repression of DNM3os
transcription.
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ATF3, a member of the ATF/CREB family of basic leucine zipper transcription factor is
an adaptive response gene. These are induced by an array of signals originating from cytokines,
physiological changes and pathologic stimuli (Filen et al., 2010; M. Gilchrist et al., 2006; T. Hai
et al., 1999; Hoetzenecker et al., 2012; Lu et al., 2006; Rynes et al., 2012; Suganami et al., 2009;
Wolford et al., 2013). We showed PlGF-mediated transcription of ATF3 was essential for
DNM3os transcriptional repression. This was demonstrated by shRNA knockdown of
endogenous ATF3, which resulted in reversal of PlGF-mediated repression, and by constitutive
ATF3 expression via exogenous ATF3 expression plasmid; this attenuated DNM3os
transcription. In silico analysis of the DNM3os promoter showed several putative cis-binding
sites for ATF3; we determined that the two sites nearest the transcription start site (within 1 kb)
of DNM3os were required for PlGF-mediated repression. This was accomplished by ATF3
promoter analysis employing a heterologous luciferase reporter. This construct was repressed by
PlGF like the endogenous gene, and furthermore was repressed, in the absence of PlGF, by co-
expressing an ATF3 cDNA gene.
We showed ATF3 shRNA reversed PlGF-mediated reduction in DNM3os reporter
activity. Concomitantly, ATF3 variant 1, a full length protein, attenuated reporter luciferase
activity, while the truncated isoform (DN-ATF3), which lacks the leucine zipper dimerization
domain binding to the ATF3 site in the promoter was unresponsive to PlGF-mediated repression.
Furthermore, mutation of ATF3 binding sites in the DNM3os promoter of the heterologous
luciferase reporter rendered the promoter unresponsive to PlGF repression. This indicated that
ATF3 binding to bona fide ATF3 sites was essential for transcriptional repression. Additionally,
PlGF treatment of HMEC-1, in the presence of ATF3 shRNA, reduced binding of ATF3 to the
endogenous DNM3os promoter, as demonstrated by ChIP assay. Thus results from both
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DNM3os reporter assay and ChIP experiments showed that ATF3 acted as a PlGF-mediated
repressor of DNM3os transcription.
The nature of the ATF3 complex involved in repression of DNM3os transcription was
investigated by proteomic analysis, wherein cell lysate from HMEC-1 cells following
transfection with ATF3 expression plasmid was immunoprecipitated with antibody to ATF3.
Mass spectroscopy of immune complexes followed by data analysis revealed ~84 polypeptides
that associated either directly or indirectly with ATF3. A gene ontology classification allowed us
to focus on proteins known to be regulators of transcription. These included ATF2, JDP2, AP-1
complex, HDAC6 and HDAC7 (Table II).
Previous studies have shown that ATF3 forms homodimer and heterodimers complexes
with basic zipper leucine proteins, i.e., ATF2, JDP2, and AP-1 complex, to regulate target genes
both positively and negatively (Darlyuk-Saadon et al., 2012; Maruyama et al., 2012; Weidenfeld-
Baranboim et al., 2009). Thus we examined the association of these proteins with ATF3 as co-
regulators. Our results showed that JDP2, but not c-Jun or ATF2 were capable of associating
with ATF3 in response to PlGF-mediated repression of DNM3os.
It has been well established that acetylation of core histones is associated with chromatin
remodeling and transcriptional activation. By contrast, deacetylation of core histones is
associated with histone methylation, the latter favoring condensation of chromatin structure and
associated gene silencing (Jenuwein & Allis, 2001; Narlikar, Fan, & Kingston, 2002) Histone
deacetylases (HDACs) are involved in modifying core histones and non-histone proteins by
removal of an acetyl group from specific lysine residues (Choudhary et al., 2009). HDAC
enzymes are organized by sequence families (Kaluza et al., 2011), denoted as class I (HDACs
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1.2,3 and 8), class II (HDACs 4,5,6,7,9 and 10), class III (sirtuins), and class IV (HDAC 11).
Our study showed HDAC6 and HDAC7 were involved in PlGF–mediated repression of
DNM3os, as demonstrated by use of shRNAs for selected HDACs. This was not due to increased
expression of HDAC6 or HDAC7 mRNA by PlGF, as has been seen for hypoxia mediated
upregulation of HDAC6 (Kaluza et al., 2011). Moreover, Tubacin, a selective inhibitor of
HDAC6 (Dallavalle, Pisano, & Zunino, 2012; Haggarty, Koeller, Wong, Grozinger, & Schreiber,
2003) decreases PlGF-mediated repression of DNM3os. This effect was highly localized as
transcription of the overlapping DNM3 gene on the sense strand was unaffected.
In order to gain insight into chromatin modification events associated with ATF3
mediated DNM3os repression, we examined the DNM3os promoter by utilizing FAIRE
(formaldehyde assisted isolation of regulatory elements) methodology. This method has been
used to identify nucleosome-free, euchromatic regions of the genome (Giresi et al., 2007).
Chromatin accessibility at the ATF3 loci in the DNM3os promoter was determined by FAIRE-
qPCR analysis. Our studies showed PlGF-mediated upregulation of ATF3 levels was associated
with chromatin condensation at cis-regulatory regions of the DNM3os promoter, consistent with
repression of DNM3os transcription.
Our investigation into the nature of the repressor complex showed HDAC6 associated
with the ATF3 binding domain of the DNM3os promoter as demonstrated by ChIP assay.
Specifically, HDAC6 interacted with ATF3 sites 1 and 2 of the DNM3os promoter, as mutation
of both sites abrogated PlGF-mediated repression. Our results showed that ATF3 site in
DNM3os proximal promoter was likely involved in HDAC6 mediated chromatin remodeling and
PlGF mediated repression of DNM3os transcription.
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Results of the ENCODE project, showed the DNM3os gene, located in chromosome 1,
has extensive histone H3K27Ac acetylation marks flanking ~0.5 kb, both 5’ and 3’ of the
transcription start site of DNM3os (nt 172,113,424 to 172,114,622). We observed PlGF caused
H3K27Ac deacetylation proximal to the ATF3 sites of the DNM3os promoter and was
associated with resulting condensation of chromatin in that region. These events were consistent
with concomitant reduction of DNM3os transcription. Conversely, silencing of HDAC6 or
treatment with histone deacetylase inhibitor, Tubacin, antagonized PlGF-induced H3K27Ac
deacetylation and maintained open chromatin to allow DNM3os transcription. Transcription of
this locus results in miR-199a2 synthesis, which is maintained by Tubacin even in the presence
of PlGF. Under these conditions miR-199a2 was expressed and subsequently attenuated HIF-1α
levels leading to decreased transcription of downstream target genes, viz. ET-1.
Since we showed higher PlGF levels were associated with increased expression of ATF3
and concomitant augmented levels of ET-1, we examined the lung tissues of BK-SS sickle mice
and normal C57BL/6NJ mice. As the site of pulmonary hypertension, we expected differences in
ATF3 expression between these animals. Indeed, our studies showed pre-miR-199a2 levels were
attenuated in lungs of BK-SS mice compared to control mice and this was associated with
increased expression of ATF3 and ET-1. Thus, both in vitro and in vivo studies were consistent
in that increased levels of ATF3 were associated with reduction of miR-199a2 leading to
augmented expression of HIF-1α and its target gene ET-1.

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CHAPTER 6: Summary of Study
Pulmonary Hypertension (PHT) is a severe complication in Sickle Cell Disease (SCD)
and is highly associated with the mortality of SCD patients and manifests with increasing age
(Gladwin et al., 2004). Previous studies showed both PlGF and ET-1 are associated with the
development of PH in SCD patients (Sundaram et al., 2010) and HIF-1α is an important
upstream transcription factor in this process for regulating ET-1 expression in response to PlGF
induction in endothelial cells (Patel et al., 2008). However, in this PlGF—HIF-1α—ET-1 axis of
PHT, little is known on how the post-transcriptional regulation by miRNA regulation, is
mechanistically involved.
In Chapter 3, we showed miR-648 regulates ET-1 mRNA stability, and subsequently, its
expression in endothelial cells by direct binding to the 3’UTR of ET-1 mRNA. Moreover, our
data showed that in endothelial cells, miR-648 is co-transcribed with its host gene MICAL3 and
a transcription factor PAX5 regulates both miR-648 and MICAL3 in endothelial cells. Further,
we extend the study to SCD patients, and concluded that SCD patients have a significant higher
level of miR-648 in plasma compared to healthy subjects. This study, for the first time, showed a
novel miRNA, miRNA-648 can regulate ET-1 in endothelial cells and miR-648 can be a
promising biomarker of SCD, targeting the transcription of miR-648 or its host gene MICAL3
may provide novel insights into new therapeutic approaches for treatment to SCD.
In Chapter 4, we focused on miR-199a, which is highly associated with another important
factor in this axis, HIF-1α. Experimental results showed in endothelial cells, PlGF reduced the
endogenous level of miR-199a, which targets HIF-1α mRNA by direct binding to its 3’UTR and
concomitantly regulates expression of the downstream effector ET-1. Similar to miR-648, miR-
199a is located in its host gene Dynamin 3 opposite strand (DNM3os) and it is found to be co-
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transcribed in this transcription unit. The transcription of DNM3os was found to be subject to
PPARα regulation. By utilizing fenofibrate, a PPARα agonist, we showed the expression of miR-
199a2 and DNM3os can be increased in endothelial cells from basal levels or during PlGF
treatment. In vivo studies of fenofibrate-fed Berkeley sickle mice showed increased levels of
plasma and lung miR-199a2 and reduced levels of ET-1 in lung tissues. This study revealed the
extensively studied connection between miR-199a and HIF-1α in cancer is also important in
PHT. The co-transcription of miR-199a and its host gene DNM3os governs endogenous levels of
miR-199a in endothelial cells, thus it is a potential target for further study. Fenofibrate, a PPARα
agonist, also showed it was capable of inducing miR-199a2 expression to ameliorate PH by
reduction of ET-1 levels, thus demonstrating this therapeutic class as a promising approach for
treatment of PHT.
In the next chapter, we discussed how PlGF regulates miR-199a and its host gene
DNM3os in endothelial cells, which involves a transcriptional factor ATF3. Our data indicated
ATF3 is repressed in endothelial cells at an early time-point (~2 hrs) following PlGF induction.
Regulatory effects of ATF3 on DNM3os and pre-miR-199a were shown by luciferase activity
assay and by gain-and-loss of function studies, utilizing ATF3 shRNA and ATF3 overexpressing
vector approaches. Moreover, since ATF3 alone does not have repressor activity, proteomic
analysis of cell lysate derived from HMEC cells expressing ATF3 and co-immunoprecipitated
with ATF3 antibody, showed association of potential ATF3 binding partners: ATF2, JDP2,
HDAC6 and HDAC7. Further results showed HDAC6 is associated with ATF3 site in DNM3os
promoter to repress the transcription of DNM3os. Tubacin, a selective inhibitor of HDAC6, can
induce the expression of DNM3os RNA and pre-miR-199a2 RNA, and subsequently reduced
PlGF-mediated expression of HIF-1α and its target gene ET-1. In vivo studies of lung tissue
139

from Berkeley Sickle (SS) mice showed levels of ATF3 are increased in these mice, which
correlated with increased expression of ET-1, a marker of pulmonary hypertension. This study
demonstrated a new regulatory mechanism that an important transcription factor ATF3 can
repress expression of miR-199a and its host gene DNM3os in endothelial cells by recruiting the
HDACs to this target gene promoter to trigger the subsequent epigenetic change and chromatin
remodeling. In this study Tubacin, a HDAC6 inhibitor, also showed a promising effect by
inducing miR-199a2 expression which could attenuate HIF-1α and ET-1 levels induced by PlGF.
Thus, Tubacin and other HDAC inhibitors can be used as potential drugs to repress the
development of PHT in SCD.
To summarize, these studies broadened our knowledge on how post-transcriptional
regulation, especially miRNA regulation, is governing HIF-1α and ET-1 level in endothelial
cells. Thus, by targeting these miRNAs, such as miR-199a and miR-648, novel miRNA-based
therapeutic approaches may be developed and utilized to repress the development of PHT in
SCD. Moreover, some evidence indicate that independent transcription does occur in the
biogenesis of particular miRNAs (Ramalingam et al., 2014). However, in most cases reported so
far, intronic miRNAs are believed to share promoters with their host genes (Ramalingam et al.,
2014; Schanen & Li, 2011), which is indeed the same situation as with miR-199a2 and miR-648.
Finally, although epigenetic changes are widely studied in diseases such as cancer there is a
dearth of literature reporting the association between epigenetics and PHT/SCD. The
ATF/HDAC6 results provide a new aspect for examining the causes and treatment of PHT in
SCD: the chromatin remodeling events are also playing a very important role in the post-
transcriptional regulation of key gene expression thus contributing to PHT and SCD. More work
is therefore expected to emerge in this field in the future.
140

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Abstract (if available)

Abstract Pulmonary hypertension (PHT) is a highly prevalent complication of Sickle Cell Disease (SCD), and it is a major cause of early morbidity and mortality in sickle cell patients. SCD patients with PHT show increased levels of Placenta Growth Factor (PlGF) and a potent vaso-constrictor Endothelin-1 (ET-1) in the plasma. The previous work from our laboratory has shown that PlGF, elaborated from erythroid cells, show high levels in the plasma of SCD patients compared to healthy subjects. PlGF has been shown to induce the expression of inflammatory cytochemokines and ET-1 through a mechanism involving the activation of hypoxia induced factor-1α (HIF-1α), independent of hypoxia. Moreover, studies showed that binding of HIF-1α to the hypoxia response elements (HRE) in the promoter region of the ET-1 gene leads to increased synthesis of ET-1 mRNA. ❧ However, in the PlGF—HIF-1α—ET-1 axis of PHT, the mechanisms by which PlGF enhances the stability of ET-1 and HIF-1α mRNA, and how HIF-1α regulates its down-stream effector ET-1 remain unclear. In Chapter 3, we showed that in human endothelial cells, the stability of ET-1 mRNA was increased in response to PlGF treatment via regulation of miR-648, which decreases ET-1 expression by direct binding to the 3’ untranslated region (3’UTR) of ET-1 mRNA. Moreover, our data showed that in endothelial cells, miR-648 is co-transcribed with Microtubule-associated monooxygenase, calponin and LIM domain containing 3 (MICAL3)

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Creator Li, Chen (author)

Core Title The role of miRNA and its regulation in pulmonary hypertension in sickle cell disease

School Keck School of Medicine

Degree Doctor of Philosophy

Degree Program Genetic, Molecular and Cellular Biology

Publication Date 10/20/2017

Defense Date 10/19/2015

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

Tag miRNA,OAI-PMH Harvest,pulmonary hypertension,sickle cell disease,transcription regulation

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Language English

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Advisor Tahara, Stanley (committee chair), Kalra, Vijay (committee member), Lu, Wange (committee member)

Creator Email 2009lichen@gmail.com,cli7@usc.edu

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