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Placenta growth factor-miRNAs-lncRNAs axis in the regulation of ET-1 gene involved in pulmonary hypertension in sickle cell disease

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Placenta growth factor-miRNAs-lncRNAs axis in the regulation of ET-1 gene involved in pulmonary hypertension in sickle cell disease

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Placenta growth factor-miRNAs-lncRNAs axis in
the regulation of ET-1 gene involved in pulmonary
hypertension in sickle cell disease
by

Ke Peng

A Thesis Presented to the
FACULTY OF THE KECK SCHOOL OF MEDICINE
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
MASTER OF SCIENCE
(BIOCHEMISTRY AND MOLECULAR BIOLOGY)

MAY 2018

Copyright 2018 Ke Peng

ii

Acknowledgements

I would like to express my sincere gratitude to my PI, Dr. Vijay Kalra, who
guided me with my project for the past two years. It was a hard but meaningful
period since I had limited biology background as an undergraduate, and the
project and thesis would not be worked out without his help and words of
encouragement. I would also like to express my sincere gratitude to my committee
members, Dr. Stanley Tahara and Dr. Zoltan Tokes, who gave me inspiration and
wonderful feedback to help me improve my thesis. I thank Dr. Tahara for guidance,
the access to the required materials and equipment for the project. I sincerely
thank them for their time and effort to help me complete this thesis.
I would also like to thank my lab members, Dr. Shuxiao Zhang and
Emiliano Huesca, for providing me the training and experiment skills. I am
particularly thankful to Dr. Zhang, who guided me during weekly lab meetings.
Additionally, I thank the senior graduate students, Kyueon Park and Xinyan Liang,
who also gave me suggestion and helped with my experiments and encouraged
me when I failed to have a good result.
Finally, I would like to express my eternal gratitude to my parents, Dr. Yu
Peng and Li Wang, for their love and support in my education and daily life.

iii

Table of Contents

Acknowledgements .............................................................................................. ii
Abstract ............................................................................................................... vii
Chapter 1: Introduction ....................................................................................... 1
1.1 Pathobiology of sickle cell disease ................................................................. 1
1.2 Pulmonary hypertension in SCD .................................................................... 2
1.3 PlGF contributes to elevated levels of ET-1 and PAI-1, which contribute to PH
.................................................................................................................. ….5
1.4 Role of miRNAs in the stabilization of PlGF mediated HIF-1α expression ...... 7
1.5 Long non-coding RNAs (lncRNAs) in the regulation of ET-1 .......................... 8
Chapter 2: Hypothesis and Specific Aims ....................................................... 12
2.1 Hypothesis ................................................................................................... 12
2.2 Specific Aims ............................................................................................... 12
Chapter 3: Materials and Methods .................................................................... 13
3.1 Cell culture................................................................................................... 13
3.2 RNA extraction and mRNA analysis............................................................. 13
3.3 Transient transfection .................................................................................. 14
3.4 Western Blots .............................................................................................. 14
3.5 ELISA assay for quantitation of ET-1. .......................................................... 15
Chapter 4: Results ............................................................................................. 18
4.1 PlGF affects ET-1 and ET-1AS mRNA levels both in t-HMEC and t-HBEC .. 18
4.2 PlGF affects ET-1 protein levels both in t-HMEC and t-HBEC ...................... 19
4.3 Effect of ET-1AS expression plasmid on ET-1 mRNA levels in t-HMEC and
t-HBEC ........................................................................................................ 20
4.4 Effect of ET-1AS expression plasmid on the ET-1 protein levels in t-HMEC
and t-HBEC. ................................................................................................ 22
4.5 Effect of ET-1AS knockdown (siRNA) on ET-1 protein expression in t-HBEC
cells ............................................................................................................. 24
4.6 Effect of ET-1AS expression and knockdown (siRNA) on the released ET-1
protein in t-HBEC cells ................................................................................. 25
Chapter 5: Discussion ....................................................................................... 26
iv

5.1 Discussion ................................................................................................... 26
5.2 Future Direction ........................................................................................... 29
5.3 Conclusions ................................................................................................. 30
Bibliography ....................................................................................................... 32

v

Table of Figures

Figure 1. Schematic progression of pulmonary hypertension in sickle cell disease
..................................................................................................................... 4
Figure 2. PlGF produced from erythroid cells activates HIF-1α in both monocytes
and endothelial cells and concomitantly increases ET-1 and PAI-1 expression..6
Figure 3. Schematic of miR-199a2/miR-214 cluster in DNM3os and transcriptional
regulation of DNM3os. ............................................................................................ 7
Figure 4. Schematic of Natural Antisense Transcripts (NATs). ........................... 9
Figure 5. Schematic of overlapping region of ET-1 and ET-1AS in human and
mouse. ................................................................................................................... 11
Figure 6. ET-1 mRNA expression in response to PlGF in t-HMEC and t-HBEC..19
Figure 7. ET-1 protein expression induced by PlGF in t-HMEC and t-HBEC. .... 20
Figure 8. Effect of ET-1AS expression in endothelial cells on expression of ET-1
mRNA. .................................................................................................................... 22
Figure 9. Effect of ET-1AS expression in endothelial cells levels...................... 23
Figure 10. Effect of expression of ET-1AS and knockdown of ET-1AS in t-HBEC
cells on the release of ET-1 protein. .................................................................... 24
Figure 11. Effect of expression of ET-1AS and knockdown of ET-1AS in t-HBEC
cells on the release of ET-1 protein. .................................................................... 25
Figure 12. Alternative-splicing model .................................................................. 28
Figure 13. Different ET-1AS deletion constructs ................................................. 30
vi

List of Tables

Table 1. List of primer sequences ........................................................................ 17
vii

Abstract

Sickle cell disease (SCD), an inherited chronic hemoglobinopathy, is one of the
most common monogenic disorders worldwide. In SCD, the hemoglobin β gene
contains a valine residue in place of glutamic acid. Under low oxygen tension the
HbS undergoes polymerization leading to the sickle shape of red blood cells (RBC)
which adhere to vascular endothelium, leading to vaso-occlusion.
Pulmonary hypertension (PH) is the leading cause of morbidity and mortality in
adult SCD patients. The central risk factor for the development of PH in patients
with hemoglobinopathies is hemolytic anemia, which induces expression of
endothelin-1 (ET-1), a vasoconstrictor leading to PH. Our previous study shows
high circulating levels of placenta growth factor (PlGF) are associated with high
levels of ET-1 in Berkeley sickle mice (BKSS) and high pulmonary artery pressure.
Normal mice (C57) injected with the adenoviral vector of PlGF also show high
circulating levels of PlGF and ET-1. Furthermore, PlGF augments the expression
of HIF-1α, independent of hypoxia. Studies from our laboratory show miR-199a2
targets the 3’UTR of HIF-1α mRNA. Since PlGF reduces the expression of
miR-199a2, the expression of HIF-1α mRNA and protein goes up and
concomitantly the expression of ET-1 increases.
Other modulators of gene expression are the long non-coding RNAs (lncRNA),
which are not translated into proteins. The genome database shows the presence
of a natural antisense transcript (NAT) for ET-1, designated as ET-1AS, which
viii

overlaps ET-1 in a tail-to-tail fashion. We hypothesize ET-1AS regulates the
expression of ET-1. Our studies showed PlGF increases RNA expression of both
ET-1AS and ET-1. Furthermore, transfection of human endothelial cell lines with
ET-1AS expression plasmid increased ET-1 mRNA levels and decreased ET-1
protein expression. Conversely, transfection with ET-1AS siRNA increased
secreted ET-1 protein expression. These results showed ET-1AS has a positive
(concordant) effect on ET-1 mRNA expression. Interestingly, ET-1AS expression
reduced ET-1 protein, while ET-1AS siRNA increased secreted ET-1 protein
suggesting that ET-1AS may affect a post-transcriptional process.

1

Chapter 1: Introduction

1.1 Pathobiology of sickle cell disease

Sickle cell disease (SCD), an inherited chronic hemoglobinopathy, is one of
the most common monogenetic disorders worldwide. Approximately 7% of the
world population carries these disorders and up to 400,000 children with severe
hemoglobinopathies are born each year. There are ~30 million people with the
sickle cell disease worldwide (Machado, 2010).
SCD occurs when a person inherits a single-nucleotide substitution in the
β-globin gene on chromosome 11 (GTG for GAG), which results in the
replacement of a glutamic acid residue with valine in the β-globin chain of
hemoglobin molecule (termed HbS) (Stuart, 2004). The normal HbA is comprised
normal α-globin and β-globin chain, and undergoes a conformational change
when the molecule binds or releases oxygen (Madigan, 2006). In SCD, HbS
undergoes polymerization at low oxygen tension (hypoxia) leading to the
formation of fibers inside the red blood cells. As a result, the shape of RBCs is
altered, i.e. sickled. When these cells pass through small capillaries, they undergo
hemolysis resulting in hemolytic anemia (Morris, 2008; Stuart, 2004). This sickled
morphology has been shown to block the microcirculation and gives rise to the
vaso-occlusive nature of the disease (Bunn, 1997). Additionally, the HbS
contributes to various clinical manifestations, including chronic anemia, acute
2

chest syndrome, splenic sequestration, chronic leg ulcers, stroke, pulmonary
hypertension and asthma/reactive airway disease (Kalra, 2018).

1.2 Pulmonary hypertension in SCD

Pulmonary hypertension (PH) is a lung disorder where pulmonary artery
pressure is above normal levels, affects oxygenation and right-heart function,
ultimately becoming life-threatening (Farber, 2004). PH is also one of the clinical
manifestations in SCD which is emerging as one of the leading causes of
morbidity and mortality in adult patients (Machado, 2010). Studies reported 20%
to 30% of patients with SCD have elevated pulmonary artery pressures (Machado,
2010) and a 50% 2-year mortality rate in patients with SCD and PH (Castro, 2003).
PH in SCD is associated with vasoconstriction, vascular smooth muscle
proliferation, and irregular endothelium in pulmonary arteries with associated
thrombosis. These conditions all contribute to luminal narrowing and eventual
right ventricular failure (Kato, 2007).
As the main manifestation in SCD, the pathogenesis of PH is relatively
complex (Kalra, 2018). The relationship between hemolysis and PH is important
because free hemoglobin can inactivate the intrinsic vasodilator nitric oxide (NO)
(Rother, 2005; Reiter, 2002). NO regulates basal vasodilator tone, inhibits platelet
and hemostatic activation, inhibits transcriptional expression of nuclear factor κB–
dependent adhesion molecules (ICAM-1, VCAM-1 and P-Selectin), and reduces
superoxide levels through radical–radical scavenging (Furchgott, 1980; Ignarro,
3

1987; Palmer, 1987; Panza, 1993; De Caterina, 1995). Furthermore, hemolysis
releases arginase, which depletes the substrate for NO synthesis (Morris, 2005).
4

Figure 1. Schematic progression of pulmonary hypertension in sickle cell
disease
Impaired NO bioavailability leads to chronic pulmonary vasoconstriction and
gradually elevates pulmonary artery pressures. Excess arginase continuously
consumes arginine, which is the obligate substrate for NO production. The
longstanding effect develops vascular smooth muscle hyperplasia and eventually
creates an irregular endothelium and further accelerates pulmonary artery
hypertension (Morris, 2008).

5

Epidemiologic studies suggest that the central risk factor for the development
of PH in patients with hemoglobinopathies is hemolytic anemia, which induces
endothelin-1 (ET-1) mediated responses in PH (Machado, 2010). In turn,
hemolytic anemia is associated with intravascular hemolysis (Watts, 2007). High
plasma ET-1 levels are also observed in patients with SCD both at steady state
and during vaso-occlusive pain crises (Ergul, 2004).

1.3 PlGF contributes to elevated levels of ET-1 and PAI-1, which contribute
to PH

Placenta growth factor (PlGF), a member of vascular endothelial growth factor
(VEGF), is an angiogenic factor identified in 1991 (Maglione, 1991). PlGF exists in
four isoforms, generated by alternative mRNA splicing in the human genome (Cao,
1997). Erythroid cells produce PlGF (Tordjman, 2001). Since there is an increased
erythropoiesis in SCD patients and the sickle mouse model, increased levels of
plasma PlGF were observed in SCD (Perlman, 2003). Furthermore, in Berkeley
sickle cell mouse, high circulating levels of PlGF are associated with high levels of
ET-1 and concomitant high pulmonary artery pressure; normal mice injected with
the adenoviral vector of PlGF also show high circulating levels of PlGF and ET-1
(Sundaram, 2010; Li, 2016). Moreover, SCD patients with PH have high
circulating levels of PlGF and ET-1, and the levels of PlGF and ET-1 increase as
the degree of PH goes from modest to severe (Sundaram, 2010).
PlGF has been shown to activate hypoxia-inducible factor 1α (HIF-1α),
6

independent of hypoxia (Patel, 2008; Patel, 2009). HIF-1α dimerizes with HIF-1β
in the nucleus, and this transcription factor has been shown to regulate the
transcription of more than 100 genes essential for hypoxic adaptation,
angiogenesis, invasion and tumor progression (Bertozzi, 2011; Darnell, 2010;
Demaria, 2010; Semenza, 1992). PlGF activation of HIF-1α subsequently
up-regulates the expression of ET-1 and PAI-1 in cultured pulmonary endothelial
cells and in vivo (Patel, 2008; Patel, 2010; Kalra, 2018). Comparision of wild-type
BALB/c controls (PlGF+/+) mice to BALB/c animals genetically lacking PlGF
(PlGF–/–) mice shows the latter have reduced ET-1 and PAI-1 levels (Sundaram,
2010; Patel, 2008; Patel, 2010; Kalra 2018).

Figure 2. PlGF produced from erythroid cells activates HIF-1α in both
monocytes and endothelial cells and concomitantly increases ET-1 and
PAI-1 expression.
Due to increased generation of erythropoietin in SCD, production of PlGF from
erythroid cells increases. PlGF then interacts with either monocytes or endothelial
cells to activate HIF-1α, independent of hypoxia. Subsequently, HIF-1α
upregulates downstream target genes i.e. ET-1 and PAI-1, both involved in
pulmonary hypertension in SCD.

7

1.4 Role of miRNAs in the stabilization of PlGF mediated HIF-1α expression

Since PlGF stabilizes HIF-1α, in the absence of hypoxia, it was hypothesized
that miRNAs may affect the expression of HIF-1α. Studies show miR-199a2
targets the 3’-UTR of HIF-1α mRNA (Li, 2014). Furthermore, miR-199a2/miR-214
is located on the opposite strand of DNM3, referred to as DNM3os, and the
expression of DNM3os is regulated by transcription factor PPAR-α (Li, 2014;
Figure 3).

Figure 3. Schematic of miR-199a2/miR-214 cluster in DNM3os and
transcriptional regulation of DNM3os.
PlGF causes decreased levels of miR-199a2/miR-214, which results in activation
of PlGF and HIF-1α, and increasing ET-1. PlGF represses DNM3os. Furthermore,
PPAR-α regulates expression of DNM3os. Treatment with Fenofibrate, a PPAR-α
agonist, increases transcription of DNM3os and miR-199a2/miR-214 cluster. As a
result, the expression of PlGF and HIF-1α goes down.
8

In PlGF treated endothelial cells the expression of miR-199a2 and miR-214 is
reduced, which results in increased HIF-1α and ET-1 (Li, 2014). These results
were validated in the BKSS mouse model. The lung tissues of BKSS mice show
significantly reduced expression of miR-199a2/miR-214, and increased levels of
ET-1 and PlGF. (Li, 2014; Kalra, 2018). These studies show the role of
miR-199a2/miR-214 in the regulation of HIF-1α and PlGF genes, and their effects
on ET-1 gene involved in pulmonary hypertension in sickle cell disease. Other
modulators of gene expression are the long non-coding RNAs (lncRNA), which
are not translated into proteins.

1.5 Long non-coding RNAs (lncRNAs) in the regulation of ET-1

Long non-coding RNAs (lncRNAs) are transcripts longer than 200 nucleotides
and do not have protein-coding potential (Qin, 2016). LncRNAs can be further
divided into natural antisense transcripts (NATs), long-intergenic non-coding
RNAs (lincRNAs) and pseudogenes according to their genomic location (Derrien,
2012). NATs are defined as non-coding RNA transcribed from the opposite DNA
strand of a coding gene, which can affect the expression of their sense gene pair
(Khorkova, 2014). Although miRNAs are also capable of regulating the expression
of genes, there are concerns about the therapies based on the miRNAs because
of their off-target effects (Kalra, 2018). However, NATs are highly specific to the
complementary gene locus, for which they can potentially provide a highly
efficient therapy method (Wahlestedt, 2013).
9

NATs are known to overlap introns, exons, promoters, enhancers, UTRs and
flanking sequences of the partner coding genes, in all combinations (Khorkova,
2014). Three configurations of NATs corresponding to different portions of the
sense-antisense (S/AS) pairs have been observed, which are head-to-head,
tail-to-tail, and fully overlapping with the coding gene (Katayama, 2005, Figure 2).
For example, WDR83 and deoxyhypusine synthase (DHPS) are shown to form a
strong tail-to-tail paring pattern (Su, 2012), and the insulin-like growth factor 1
receptor (IGF1R) shows a very strong antisense CAGE tag overlapping the
promoter of the sense transcript (Sleutels, 2002).

There are multiple ways NATs can affect the expression of genes. First, NATs
can affect the chromatin. LncRNA may associate with accessory factors and
Figure 4. Schematic of Natural Antisense Transcripts (NATs).
Antisense transcripts can have overlapping strand with sense strand. Here are (a)
head-to-head, (b) and (d) overlapping and (c) tail-to-tail representation (Katayama,
2005).

10

direct the complex to particular target genes through both base pairing to DNA
and secondary–tertiary structure-driven protein binding (Davidovich, 2013).
Recent studies show that the IL-1β antisense transcript can change the chromatin
structure of the IL-1β promoter by decreasing H3K4 trimethylation on the
promoter, which inhibits the transcription of IL-1β (Lu, 2014). Second, NATs can
affect the post-transcription process. It is reported that the cis S/AS pairing
between WDR83 and DHPS can form a sense-antisense RNA duplex at their
overlapping regions, which might reduce mRNA stability (Su, 2012). Additionally, it
is reported that lncRNA is involved in nonsense-mediated mRNA decay (NMD),
which surveys newly synthesized mRNAs and degrades those that harbor a
premature termination codon (PTC), thereby preventing the production of
truncated proteins (Kurosaki, 2016). Recent studies show that the pre-mRNA
splicing deposits a mark named EJC about 20-24 nucleotides upstream of 80%
exon-exon junctions, which triggers NMD (Le Hir, 2000; Le Hir, 2001; Sauliere,
2012; Singh, 2012). Studies also show that a single 3’ UTR EJC deposited at an
exon-exon junction residing more than about 50-55 nucleotides downstream of
the PTC can interact with the up-frameshift protein (UPF1) complex (Gehring,
2003; Hwang, 2010; Kim, 2001; Shibuya, 2004; Yamashita, 2009). UPF1 is the
key human NMD factor whose phosphorylation represses any additional
translation initiation events on NMD targets, which is important for mRNA decay
(Isken, 2008). Recent analyses reveal that the hypophosphorylated UPF1
associates with regions of cellular RNAs that are physically accessible to its
11

binding, including lncRNAs (Gregersen, 2014; Hogg and Goff, 2010; Hurt, 2013;
Kurosaki and Maquat, 2013; Lee, 2015; Zünd, 2013).
Studies also report that antisense RNA can interact with sense mRNA
stabilization. The antisense strand of Apolipoprotein A-IV (APOA4), APOA4-AS,
regulates the expression of APOA4 at both mRNA and protein levels. Recent
investigation shows APOA4-AS has two putative HuR (mRNA stablilizing protein)
binding sites, which indicates APOA4-AS may recruit HuR to form a complex to
stabilize APOA4 mRNA (Qin, 2016). In conclusion, NATs are highly involved in the
regulation of gene expression. Thus, we hypothesize that the antisense of ET-1,
which is ET-1AS, can also affect the ET-1 expression (Figure 4).

Figure 5. Schematic of overlapping region of ET-1 and ET-1AS in human
and mouse.
As shown ET-1AS overlaps ET-1 in a tail-to-tail configuration in human, where
the exon 4 of ET-1AS overlaps with the exon 4-intron-exon 5 of ET-1. In mouse,
ET-1AS overlaps exon 2-3 and portions of introns 1 and 2.
.

12

Chapter 2: Hypothesis and Specific Aims

2.1 Hypothesis
The aim of this study is to determine the mechanism by which ET-1AS affects
the expression of ET-1 under basal and PlGF mediated conditions, the latter is
relevant to SCD. Preliminary studies show PlGF augments expression of ET-1
and ET-1AS in a time dependent manner (0.5-24 h) in the transformed human
dermal microvascular endothelial cell line (t-HMEC) and transformed human brain
endothelial cell line (t-HBEC). Furthermore, ET-1AS siRNA significantly reduced
ET-1 mRNA expression. Thus, we hypothesize ET-1AS modulates ET-1
expression, though the mechanism by which it occurs remains to be elucidated.
The detailed process of this NAT-involved ET-1 regulation will provide insight to
both NAT-involved regulation and clinical therapy to PH in SCD patients.
2.2 Specific Aims
Since elevated ET-1 levels are associated with endothelial cells dysfunction,
human dermal microvascular endothelial cell line and human brain endothelial cell
line will be used as model systems in this study. We will determine whether
ET-1AS alters the expression of ET-1 with or without PlGF. Then we will examine
which region of ET-1AS affects the regulation. Lastly, we will investigate the
mechanism by which ET-1As regulates ET-1 expression at both mRNA and
protein levels.
13

Chapter3: Materials and Methods

3.1 Cell culture
t-HMEC (transformed human dermal mictrovascular endothelial cell) and
t-HBEC (transformed human brain microvascular endothelial cells) were cultured
in RPMI-1640 containing 10% fetal bovine serum. The medium was changed
every 2 to 3 days. HMEC and HBEC (4 x 10
6
cells) were incubated in serum free
RPMI for 3 h prior to PlGF (250 ng/uL) treatment.

3.2 RNA extraction and mRNA analysis
Ribozol (Amresco, Solon, OH) was used to extract total RNA. 100 ng of RNA
was used as a template for real-time reverse transcription polymerase chain
reaction (qRT-PCR). qRT-PCR was executed using iScript One-Step RT-PCR Kit
(Bio-Rad) to quantify the mRNA expression. All primers used in mRNA analysis
are listed in Table 1. qRT-PCR was carried out following the listed steps: cDNA
synthesis at 42°C for 5 min, reverse transcriptase inactivation at 95°C for 10 s,
followed by 40 cycles of denaturation at 95°C for 5 s and annealing at 60°C for 34
s. A reference gene, GAPDH expression was used to normalize the mRNA
expression of the samples. Relative mRNA expressions were calculated as
2
− ∆ ∆ 𝐶𝑡
, where ∆ ∆ Ct = (Ct target gene of treated sample – Ct reference gene of
treated sample) – (Ct target gene of control sample – Ct reference gene of control
sample).
14

3.3 Transient transfection
t-HMEC and t-HBEC cells (1 x 10
6
cells) were transfected with an ET-1AS
expression plasmid (1 µg) and siRNA against ET-1AS (1.5 µM) suspended in PBS.
The plasmids were added to 100 µL of cell suspension followed by electroporation
using AMAXA Biosystems NucleofectorTM II. The cells were grown in a CO2
incubator for 24 h at 37°C, and then lysed in Ribozol and RIPA buffer for mRNA
and protein analysis, respectively.

3.4 Western Blots
t-HMEC and t-HBEC cells (1 x 10
6
cells), as indicated, were either treated
with PlGF or transfected by electroporation with plasmids, followed by a wash with
PBS and lysis in RIPA buffer for 30 min at 4°C. The lysates were centrifuged at
10,000 g for 1 min at 4°C and the supernatant was collected. Bradford protein
assay was performed to determine protein concentrations, and approximately 25
µg of each sample was mixed with Laemmli sample buffer for Western Blotting.
The samples were heated at 95°C for 10 min and loaded onto SDS-PAGE gels.
The proteins were separated by electrophoresis for 2 h at 100 V and transferred
electrophoretically to PVDF for 90 min. PVDF was blocked with 5% milk-TBST
solution for 1 h at room temperature prior to incubation with the diluted primary
antibody (1:400) at 4°C overnight. The PVDF membrane was washed three times
15

for 10 min in 1% TBST and incubated with secondary antibody (HRP-conjugated
rabbit antibody (1:1000) for 30 min at room temperature. The membrane was
washed three times for 10 min. Pierce ECL Western Blotting Substrate kit was
used to develop and detect HRP activity of the membrane. GAPDH was used as a
loading control and quantification of bands on the film was performed using the
image software system.

3.5 ELISA assay for quantitation of ET-1.
t-HBEC cells (106 cells) suspended in PBS were transfected with either
ET-1AS expression plasmid (1 µg) or siRNA against ET-1AS (1.5 µM). The
plasmids were added to 100 µL of cell suspension followed by electroporation
using AMAXA Biosystems NucleofectorTM II. The cells were grown in a CO2
incubator for 24 h at 37°C, and then starved in 2 mL of serum-free medium for 3 h.
After that, HBEC cells are treated with PlGF (250 ng/mL) for 16 h. The
supernatants were assayed for ET-1 release using an enzyme-linked
immunosorbent assay (ELISA) kit (R&D Systems, Minneapolis, MN). The
standard curve was generated utilizing endothelin-1 standards (0.0 pg/mL, 0.39
pg/mL, 0.78 pg/mL, and 1.56 pg/mL). 150 µL of diluted calibrator were added to
each well and mixed with 75 µL of either working standards or samples. The
ELISA plate was then covered and incubated for 1 h at room temperature on a
horizontal orbital microplate shaker set at 500 rpm. Each well was then washed
four times with wash buffer (400 µL) using Nunc-ImmunoTM Wash 8 channel.
16

Then 200 µL of Endothelin-1 monoclonal antibody conjugated to HRP was added
to each well, followed by incubation of the plate on the shaker for 3 h. Each well
was washed again four times with wash buffer (400 µL). After that, 200 µL of
substrate solution (Hydrogen peroxide and tetramethylbenzidine, 1:1) was added
to each well. The plate was incubated 30 minutes at room temperature, covered
to protect from light. Finally, 50 µL of stop solution (2 N Sulfuric acid) was added to
each well. The optical density was determined using a microplate reader
(FLUOstar Omega, BMG LABTECH). The reading was first set to 450 nm and
then 570 nm for correction. The OD at 450 nm is subtracted from OD at 570 nm
for correction. This subtraction will correct for imperfections in the plate curvature.
All measurements are assayed in duplicate. The OD is converted to pg/ml utilizing
the standard curve.

17

Table 1. List of primer sequences

Organism/
Gene
Method Forward Primer (5’→ 3’) Reverse Primer (5’→ 3’)
H. GAPDH qPCR AACCTGCCAAGTACG
ATGACATC
GTAGCCCAGGATGCC
CTT GA
H. ET-1
exon 1&2
qPCR GAAACCCACTCCCAGT
CCAC
CGGGAGTGTTGACCCAA
ATG
H. ET-1AS
exon 1&2
qPCR CACATCTGATGCCCAG
TCC
GACGGCTAACATCTGAG
TCC
H.
ET-1AS
exon 4&5
qPCR GCTGGAGCCCTCTGTG
TTC
CGACATTTCAGGGAGAA
ACTC
18

Chapter 4: Results

4.1 PlGF affects ET-1 and ET-1AS mRNA levels both in t-HMEC and t-HBEC

The previous study from our lab showed PlGF induces ET-1 expression in
cultured endothelial cells (Patel, 2008). Here we examined the effect of PlGF on
ET-1 and ET-1AS mRNA expression. We utilized two endothelial cell lines, SV-40
transformed human dermal microvascular endothelial cells (t-HMEC) (Yeligar,
2009) and SV-40 transformed human brain endothelial cells (t-HBEC) (Gonsalves
and Kalra, 2010)
Treatment of t-HMEC with PlGF showed a time-dependent change in the
expression of ET-1 mRNA and ET-1AS RNA (Figure 6A). There was a 6-fold
increase in ET-1 mRNA at 4 h, while ET-1AS increased by ~16-fold at the same
time point. Similarly, PlGF augmented the expression of ET-1 mRNA and ET-1AS
RNA in t-HBEC. Here, the effect was most pronounced at 16 h. There was a 3-fold
increase in ET-1 mRNA and also a 3-fold increase in ET-1AS RNA expression
(Figure 6B). We utilized exon 1-2 primers as well as exon 4-5 primers for
quantitation of ET-1AS RNA by qRT-PCR. The exon1-2 primers were more
effective compared to exon 4-5 primers.
19

4.2 PlGF affects ET-1 protein levels both in t-HMEC and t-HBEC

Next, we examined whether PlGF affects ET-1 protein expression. Both
t-HMEC and t-HBEC were treated with PlGF for various time periods (0.5-24 h).
The cell lysate was prepared followed by Western blot analysis utilizing rabbit
anti-ET-1 (1:400 dilution) followed by anti-rabbit-HRP. As shown in Figure 7 (A
and B) PlGF induced ET-1 protein expression in both t-HBEC and t-HMEC in a
time dependent manner (0.5-24 h). There was a 4-fold increase in ET-1 protein
expression in t-HMEC at 8 h, while a ~4.5-fold increase in ET-1 protein was
Figure 6. ET-1 mRNA expression in response to PlGF in t-HMEC and
t-HBEC.
t-HMEC and t-HBEC cells were starved in serum free media for 3 h followed by
treatment with PlGF (250 ng/ml) for 0.5, 2, 4, 8, 16 and 24 h. RNA was
extracted followed by qRT-PCR utilizing primers for ET-1, ET-1AS (exon 1-2),
ET-1AS (exon 4-5), and GAPDH. The values were normalized to GAPDH.
Data represent the mean ±SEM of nine biological replicates in t-HMEC and six
biological replicates in t-HBEC.

20

observed at 16 h in t-HBEC (Figure 7B).

4.3 Effect of ET-1AS expression plasmid on ET-1 mRNA levels in t-HMEC
and t-HBEC

Our study showed PlGF induced both ET-1 and ET-1AS RNA and ET-1
protein levels. Next, we examined the effect of ET-1AS on ET-1 mRNA and
protein in the absence of PlGF. Thus, we overexpressed ET-1AS in both t-HMEC
Figure 7. ET-1 protein expression induced by PlGF in t-HMEC and t-HBEC.
t-HMEC and t-HBEC cells were treated with PlGF (250 ng/ml) for 0.5, 2, 4, 8, 16 and
24h. Cell lysates were prepared followed by Western blot analysis utilizing antibody
to ET-1 and GAPDH. The GAPDH antibody was used to correct for loading control in
each lane. The blots were scanned and analyzed by Image J software. Data
represent the mean ±SEM of six biological replicates.
21

and t-HBEC cells without PlGF treatment and performed qRT-PCR to assess
ET-1 mRNA levels. First, we determined whether transfection with ET-1AS led to
an increase in ET-1AS RNA. As shown in Figure 8A, t-HMEC cells transfected
with ET-1AS plasmid for 24 h resulted in a ~8-fold increase in ET-1AS RNA
compared to GFP control as determined by qRT-PCR. However, there was no
significant (~1.2 fold) change in ET-1 mRNA levels (Figure 8A and B). Similar
results were observed in response to transfection of the ET-1AS plasmid in
t-HBEC cells. However, there was several hundred-fold increase in ET-1AS
(~900-fold) in t-HBEC, though there was no change in ET-1 mRNA levels (Figure
8C and D).
22

4.4 Effect of ET-1AS expression plasmid on the ET-1 protein levels in
t-HMEC and t-HBEC.

Next, we examined the effect of ET-1AS transfection of endothelial cells on
ET-1 protein expression. Cell lysates of ET-1AS transfected cells were examined
for ET-1 protein expression by Western Blot analysis. As shown in Figure 9A,
Figure 8. Effect of ET-1AS expression in endothelial cells on expression of
ET-1 mRNA.
t-HMEC and t-HBEC cells (1x10
6
cells) were electroporated with ET-1AS
expression plasmid (1 µg) and cells allowed to grow for 24 h. RNA was extracted
and qRT-PCR was performed utilizing primers for ET-1AS (exon 1-2 or exon 4-5),
ET-1 and GAPDH. Data represent the mean ±SEM of three biological replicates in
t-HMEC and two biological replicates in t-HBEC.

23

Figure 9. Effect of ET-1AS expression in endothelial cells levels.
t-HMEC and t-HBEC cells were electroporated with ET-1AS expression plasmid
(1µg) and cells allowed to grow for 24 h. Cell lysates were prepared for Western
Blot analysis of ET-1 protein. The gels were scanned for quantitation of ET-1
protein using GAPDH as a loading control. Data represent the mean ±SEM of five
biological replicates in t-HMEC (A) and three biological replicates in t-HBEC (B).
ET-1 protein levels in t-HMEC slightly increased (~50%) compared with
non-treated (control), and showed ~20% increase when compared with GFP
control. In t-HBEC, the ET-1 protein levels decreased ~ 60% compared with
non-treated (control), and showed a 90% decrease compared to GFP control
(Figure 9B). It appears that the effect of ET-1AS on ET-1 protein levels is
cell-specific.

24

4.5 Effect of ET-1AS knockdown (siRNA) on ET-1 protein expression in
t-HBEC cells

We examined the effect of ET-1AS knockdown with siRNA on the ET-1 protein
expression using Western Blot. Cell lysates of ET-1AS siRNA transfected cells
were examined for ET-1 protein expression. As shown in Figure 10, knockdown of
ET-1AS showed a ~3.5-fold increase in ET-1 protein expression compared to
non-treated (control), and a ~3-fold increase compared to scramble siRNA.

Figure 10. Effect of expression of ET-1AS and knockdown of ET-1AS in
t-HBEC cells on the release of ET-1 protein.
t-HBEC cells were electroporated for transfected of siRNA against ET-1AS (1.5 µM)
and scramble siRNA as control and allowed to grow for 24 h. Cell lysates were
prepared for Western Blot analysis of ET-1 protein. The gels were scanned for
quantitation of ET-1 protein using GAPDH as a loading control. The blot was
spliced because of other unrelated samples loaded in the same blot. Data
represent the mean ±SEM of two biological replicates.
25

4.6 Effect of ET-1AS expression and knockdown (siRNA) on the released
ET-1 protein in t-HBEC cells

Finally, we examined the effect of both ET-1AS expression and knockdown on
the secreted ET-1 protein using ELISA assay. As shown in Figure 11,
overexpression of ET-1AS did not significantly affect the release of ET-1 protein
compared to GFP control. However, knockdown of ET-1AS (ET-1AS siRNA)
showed an increase in ET-1 protein release compared to transfection with
scrambled ET-1AS. These data showed ET-1AS knockdown increased secreted
ET-1 protein levels while the overexpression of ET-1AS did not alter secreted
ET-1 protein levels.

0
0.2
0.4
0.6
0.8
1
1.2
ctrl siET1AS siSCRB ET1ASO/E GFP
Relative levels of ET-1 protein secreted
Figure 11. Effect of expression of ET-1AS and knockdown of ET-1AS in
t-HBEC cells on the release of ET-1 protein.
t-HBEC cells were electroporated and transfected with siRNA against 5’ of ET-1AS
(1.5 µM) and ET-1AS overexpression plasmid (1 µg). Supernatants were harvested
after 24 h for ELISA assay. Data represent the mean ±SEM of three biological
replicates.

26

Chapter 5: Discussion

5.1 Discussion

Patients with sickle cell disease (SCD) develop hemolytic anemia. There is an
increased destruction of red blood cells, which results in the increased presence
of hemoglobin and its oxidation product, bilirubin and hemin in the circulation.
Hemin has been shown to upregulate by several-fold the expression of placenta
growth factor in erythroid cells (Wang, 2014). Previous studies by Kalra and
coworkers show plasma levels of PlGF are significantly higher compared to
healthy individuals (Kalra, 2018). This occurs due to increased erythropoiesis in
SCD leading to the increased presence of erythroblast cells, which produce PlGF
in response to erythropoietin. Furthermore, PlGF has been shown to activate
HIF-1α, independent of hypoxia in cultured endothelial cells. Studies show
miR-199a2 targets the 3’UTR of HIF-1α mRNA. In the BKSS mouse models, the
levels of miR-199a2 are significantly reduced compared to C57 control mice (Li,
2014). When the levels of miR-199a2 decline, the HIF-1α levels will increase and
thus ET-1 goes up.

The genomic location of miR-199a2 is in the opposite strand of DNM3 i.e.
DNM3os. It has been shown PlGF down regulates the expression of DNM3os in
cultured endothelial cells (Li, 2016). As a result of miR-199a2 decrease, HIF-1α
increases. These studies show the role of miRNAs in the regulation of PlGF
27

mediated HIF-1α expression (Kalra 2018). Next, we focused our studies on long
non-coding RNAs. Specifically, we examined the role of natural antisense
transcripts (NATs) as these have been shown to affect the sense genes
(Katayama 2005). Although several thousand (15,000-55,000) NATs have been
identified in the genome, a very small number of NATs have been functionally
characterized (Qin 2016). It has been shown that APOA4-AS regulates the
expression of APOA4 lipoprotein (Qin 2016); knockdown of APOA4-AS reduced
APOA4 expression both in vitro and in vivo (Qin 2016).

Our present studies show that cultured endothelial cells, transformed human
dermal microvascular endothelial cell line (t-HMEC) and transformed human brain
endothelial cell line (t-HBEC) express ET-1AS transcript as determined by
qRT-PCR. Moreover, PlGF increases the expression of ET-1AS in a time
dependent manner. Furthermore, as previously shown, PlGF increased ET-1
expression both at the mRNA and protein levels.

To further determine the regulatory role of ET-1AS in the expression of ET-1,
we transfected ET-1AS plasmid into t-HMEC and t-HBEC cells. We observed
ET-1AS expression did not significantly affect the ET-1 mRNA, but caused a
reduction in ET-1 protein in t-HBEC cells. Furthermore, knockdown of ET-1AS by
siRNA showed an increase in ET-1 protein release. This may occur by the effect
of ET-1AS on the splicing of ET-1 gene. Under basal condition the default splicing
28

of pre-ET-1 mRNA occurs. However, when we overexpress ET-1AS, it might
interfere with the splicing of the exon 4 and exon 5 since ET-1AS overlaps the
splice acceptor site of ET-1-exon 5 (Figure 12). Additionally, there is a poly-A site
at ~500 bp downstream of exon 5 in ET-1 pre-mature mRNA. Thus when this
alternative splicing occurs, the translation of the alternatively spliced ET-1 mRNA
would give a truncated protein because of the in-frame stop codon. Furthermore,
the alternative spliced ET-1 mRNA has been reported in vivo (Totoki, 2006).

Figure 12. Alternative-splicing model
Under basal condition, the majority of pre-mRNA of ET-1 will undergo default
splicing. Note that the 3’-ends of pre-ET-1 mRNA and ET-1AS RNA are
complementary and the last exon of ET-1AS overlaps the splice acceptor site of
ET-1-exon 5. Translation of alternatively spliced ET-1 mRNA would give a
truncated ET-1 protein due to the presence of an in-frame stop codon in the last
intron. This mRNA variant may be less active in translation, resulting in decreased
ET-1 protein levels.
29

5.2 Future Direction

This study demonstrated that PlGF induced ET-1 expression in t-HMEC and
t-HBEC cells, and ET-1AS expression decreases ET-1 protein. Knockdown of
ET-1AS in these cells increased ET-1 protein secretion. It is possible that ET-1AS
decreases the translation of ET-1 protein through alternative splicing in ET-1
mRNA. Thus, it is important to examine whether there is an increasing or
decreasing level of the alternative splicing transcript with ET-1AS overexpression
or knockdown. Next, it will be important to determine the effective region of
ET-1AS that actually plays a role in the ET-1 regulation. Since the overlapping
region between ET-1 and ET-1AS contains both intron and exon, it would be
helpful to figure out the exact region in ET-1AS that affects the regulation of ET-1.
This study requires the construction of plasmids containing different parts of
ET-1AS, which would be transfected into t-HMEC and t-HBEC respectively
(Figure 13).
Moreover, Tatsuaki and colleagues have established a nonsense-mediated
mRNA decay (NMD) model in humans (Kurosaki, 2016). In this model, NMD
surveys newly synthesized mRNAs and degrades those that harbor a premature
termination codon (PTC) with the help of the key human NMD factor,
up-frameshift protein 1 (UPF1) (Kurosaki, 2016). It is possible that ET-1AS
regulates ET-1 through activating the PTC and therefore the alternative spliced
ET-1 mRNAs are degraded. Further studies to pull down the UPF1 protein with
ET-1AS expression or knockdown would be helpful.
30

5.3 Conclusions

Pulmonary hypertension (PH) is the leading cause of morbidity and mortality
in adult SCD patients. Patients with SCD and PH have high circulating levels of
PlGF, which activates the expression of ET-1. High PlGF and ET-1 levels are
observed in Berkeley sickle cell mouse. In the present study, we demonstrated the
PlGF increased ET-1AS expression. However, ET-1AS expression did not
significantly change ET-1 mRNA levels but decreased ET-1 protein. Moreover,
knockdown of ET-1AS increased secreted ET-1 protein expression. These
studies might indicate that ET-1AS decreased translation of ET-1 protein, as a
Figure 13. Different ET-1AS deletion constructs
The 270 bp region as indicated above of ET-1AS overlaps the intron of ET-1, while
the 528 bp region overlaps the exon 5 of ET-1. Different deletion constructs of
ET-1AS will be generated to test the region that affects the regulation of ET-1,
including the deletion in 270 bp, the deletion in 528 bp, and the 270 bp region only.
31

consequence of an alternative splicing mechanism. Long non- coding RNAs have
a potential role in therapeutics as these have a direct effect on specific genes
compared to miRNAs which can have off target gene effects (Wahlestedt, 2013).

32

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Abstract (if available)

Abstract Sickle cell disease (SCD), an inherited chronic hemoglobinopathy, is one of the most common monogenic disorders worldwide. In SCD, the hemoglobin β gene contains a valine residue in place of glutamic acid. Under low oxygen tension the HbS undergoes polymerization leading to the sickle shape of red blood cells (RBC) which adhere to vascular endothelium, leading to vaso‐occlusion. ❧ Pulmonary hypertension (PH) is the leading cause of morbidity and mortality in adult SCD patients. The central risk factor for the development of PH in patients with hemoglobinopathies is hemolytic anemia, which induces expression of endothelin-1 (ET-1), a vasoconstrictor leading to PH. Our previous study shows high circulating levels of placenta growth factor (PlGF) are associated with high levels of ET-1 in Berkeley sickle mice (BKSS) and high pulmonary artery pressure. Normal mice (C57) injected with the adenoviral vector of PlGF also show high circulating levels of PlGF and ET-1. Furthermore, PlGF augments the expression of HIF-1α, independent of hypoxia. Studies from our laboratory show miR-199a2 targets the 3’UTR of HIF-1α mRNA. Since PlGF reduces the expression of miR-199a2, the expression of HIF-1α mRNA and protein goes up and concomitantly the expression of ET-1 increases. ❧ Other modulators of gene expression are the long non‐coding RNAs (lncRNA), which are not translated into proteins. The genome database shows the presence of a natural antisense transcript (NAT) for ET-1, designated as ET-1AS, which overlaps ET-1 in a tail‐to‐tail fashion. We hypothesize ET-1AS regulates the expression of ET-1. Our studies showed PlGF increases RNA expression of both ET-1AS and ET-1. Furthermore, transfection of human endothelial cell lines with ET-1AS expression plasmid increased ET-1 mRNA levels and decreased ET-1 protein expression. Conversely, transfection with ET-1AS siRNA increased secreted ET-1 protein expression. These results showed ET-1AS has a positive (concordant) effect on ET-1 mRNA expression. Interestingly, ET-1AS expression reduced ET-1 protein, while ET-1AS siRNA increased secreted ET-1 protein suggesting that ET-1AS may affect a post‐transcriptional process.

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Asset Metadata

Creator Peng, Ke (author)

Core Title Placenta growth factor-miRNAs-lncRNAs axis in the regulation of ET-1 gene involved in pulmonary hypertension in sickle cell disease

School Keck School of Medicine

Degree Master of Science

Degree Program Biochemistry and Molecular Biology

Publication Date 04/23/2018

Defense Date 03/21/2018

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

Tag Endothelin-1,gene regulation,lncRNA,miRNA,natural antisense transcript,OAI-PMH Harvest,placenta growth factor,pulmonary hypertension,sickle cell disease

Format application/pdf (imt)

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Kalra, Vijay Kumar (committee chair), Tahara, Stanley (committee member), Tokes, Zoltan (committee member)

Creator Email cocopeng198@outlook.com,kepeng@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c40-494087

Unique identifier UC11267302

Identifier etd-PengKe-6262.pdf (filename),usctheses-c40-494087 (legacy record id)

Legacy Identifier etd-PengKe-6262.pdf

Dmrecord 494087

Document Type Thesis

Format application/pdf (imt)

Rights Peng, Ke

Type texts

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

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

Repository Name University of Southern California Digital Library

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