A Dissertation
entitled
Regulation of Na/K-ATPase and its Role in Cardiac Disease
by
Xiaoming Fan
Submitted to the Graduate Faculty as partial fulfillment of the requirements for the
Doctor of Philosophy Degree in Biomedical Sciences
______Dr. Jiang Tian, Committee Chair
______Dr. Andrew D. Beavis, Committee Member
______Dr. Kevin Z. Pan, Committee Member
______Dr. David Giovannucci, Committee Member
______Dr. Lijun Liu, Committee Member
______Dr. Christopher J. Cooper, Committee Member
______Dr. Cyndee Gruden, Dean College of Graduate Studies
The University of Toledo
December 2018
Copyright 2018, Xiaoming Fan
This document is copyrighted material. Under copyright law, no parts of this document may be reproduced without the expressed permission of the author. An Abstract of
Regulation of Na/K-ATPase and its Role in Cardiac Disease by
Xiaoming Fan
Submitted to the Graduate Faculty as partial fulfillment of the requirements for the Doctor of Philosophy Degree in Biomedical Sciences
The University of Toledo
December 2018
Heart failure is an important public health issue and a leading cause of mortality in the United States. Previous publications have shown that both protein amount and enzyme activity of cardiac Na/K-ATPase were reduced in heart failure patients. Analysis of gene expression database also demonstrated that expression of Na/K-ATPase α1 subunit in heart tissue from heart failure patients is lower compared to non-heart failure patients. However, it is not clear whether Na/K-ATPase reduction is causatively related with heart failure, and if so, how Na/K-ATPase expression is regulated. During the past twenty years tremendous work have been done to show that Na/K-ATPase, especially its signaling function, is associated with cardiac hypertrophy and cardiac fibrosis. Interestingly, data from our laboratory showed that reduction of Na/K-ATPase attenuates its signaling activation, which might be beneficial to reducing cardiac hypertrophy, but it also increases cardiac cell apoptosis. The aim of my project is to: 1) study how Na/K-ATPase is regulated during disease conditions; and 2) study the effect of Na/K-ATPase reduction on cardiac function and its mechanism. From these studies, we have identified a novel long-non-coding RNA,
ATP1A1-AS1, as an endogenous Na/K-ATPase regulator that affect Na/K-ATPase expression and its signaling function (Chapter 3). We also demonstrated that reduction of
iii Na/K-ATPase α1 had significant effect on cardiac function and remodeling due to the change in Na/K-ATPase signaling (Chapter 4).
ATP1A1-AS1 gene is a natural antisense gene of Na/K-ATPase α1 (ATP1A1) which is located on the opposite strand of the Na/K-ATPase α1 gene. Our results showed that 4 splice variants expressed in human adult kidney cells (HK2 cells) and embryonic kidney cells (HEK293 cells). We found that inhibition of DNA methylation had a differential effect on the expression of ATP1A1-AS1 and its sense gene. To investigate the physiological role of this antisense gene, overexpression of ATP1A1-AS1 was performed and its effect on Na/K-ATPase expression was examined. The result showed that ATP1A1-
AS1-203 overexpression reduced the Na/K-ATPase α1 (ATP1A1) gene expression in HK2 cells by about 20% (p<0.05), it also reduced the Na/K-ATPase α1 protein by about 22%
(p<0.05). In addition, overexpression of ATP1A1-AS1-203 attenuated ouabain-induced
Src activation in HK2 cells and subsequently inhibited the cell proliferation in the presence or absence of ouabain. These results demonstrate that ATP1A1-AS1 gene is a moderate negative regulator of Na/K-ATPase α1 and can modulate Na/K-ATPase-related signaling pathways. Importantly, these results suggest that a moderate reduction of Na/K-ATPase expression could disproportionally affect the signaling function of Na/K-ATPase, which is consistent to the previous findings in Na/K-ATPase knockdown cell lines.
To study the effect of Na/K-ATPase on cardiac function in disease conditions, we used a mouse chronic kidney disease (CKD) model (5/6th partial nephrectomy or PNx) in
Na/K-ATPase alpha 1 heterozygous (α1+/-) mice and their wild type (WT) littermates. The cardiac Na/K-ATPase alpha 1 expression in α1+/- mice is about 40% less than that of WT mice. The experimental results showed that reduction of Na/K-ATPase significantly
iv reduced cardiac hypertrophy in the CKD animals. However, it showed no significant change in cardiac fibrosis between the two animal strains. To further study the role of
Na/K-ATPase in cardiac fibrosis, we found that CKD induces activation of Src and NFB signaling in the heart tissue of WT mice, which subsequently causes reduction of microRNA-29b-3p (miR-29b), an antifibrotic microRNA. However, in α1+/- mice, Src and
NFB activation was significantly attenuated. We further found that in WT mice, inhibition of Src signaling using pNaKtide blocked NFB activation and restored miR-29b expression to a level close to the controls. Whereas, the pNaKtide had no significant effect on Src/NFB activation or miR-29b expression in α1+/- mice. Injection of pNaKtide also significantly reduces cardiac fibrosis in WT mice. These results suggest that Na/K-ATPase reduction not only attenuates Na/K-ATPase signaling function, it may also adopt other pathological pathways, which can cause cardiac fibrosis independent of Na/K-ATPase- related Src and NFB signaling. The specific mechanisms are elusive and merit further investigations.
v
To my wife, Dr. Xiaolu Zhang, for your love and support during my Ph.D. study.
To my parents, Hansong Fan and Xuanfen Luo, for your love, support, and encouragement throughout my life.
To my son, Theo Fan, for the joy you brought to my life.
vi
Acknowledgements
I am extremely grateful to Dr. Jiang Tian for his guidance and faithful support throughout my graduate studies in both academic and personal life. I am looking forward to work with you in the future.
I would like to thank my academic advisory committee Dr. Andrew Beavis, Dr.
Kevin Z. Pan, Dr. David Giovannucci, Dr. Christopher J. Cooper and Dr. Lijun Liu, for their expert advice and guidance throughout my graduate studies.
I would also like to appreciate the support from the Department of medicine, especially the support from Dr. Lance Dworkin.
My thanks also go to the people in Dr. Kumarasamy’s lab, Dr. Kennedy’s lab, Dr.
Haller’s lab, Dr. Gupta’s lab, and Dr. Gong’ lab for the great collaboration and help during my graduate study.
A special thanks to Dr. Shihe Liu for his friendly help.
I would especially like to thank all my former and current colleagues Dr.
Christopher Drummond, Dr. Huilin Shi, Xie, Dr. Jeffrey Xinshuo Xie and Mano
Tillekeratne, for all the great discussion, inspiration and friendship.
vii
Table of Contents
Abstract ...... iii
Acknowledgements ...... vii
Table of Contents ...... viii
List of Tables ...... x
List of Figures ...... xi
List of Abbreviations ...... xiii
1. Introduction ...... 1
2. Literature review ...... 4
2.1 Na/K-ATPase biology ...... 4
2.1.1 Structure of Na/K-ATPase ...... 4
2.1.2 Ion pumping function of Na/K-ATPase ...... 6
2.1.3 Ligands of Na/K-ATPase ...... 7
2.1.4 Signaling function of Na/K-ATPase ...... 8
2.2 Heart failure and cardiac remodeling ...... 9
2.3 Na/K-ATPase and cardiac remodeling ...... 11
2.4 Regulation of Na/K-ATPase ...... 14
3. Characterization of A Long Non-Coding RNA, The Antisense RNA of Na/K-ATPase
Alpha 1 In Human Kidney Cells ...... 16
viii Abstract ...... 17
Introduction ...... 18
Results ...... 20
Discussion ...... 23
Materials and Methods ...... 25
Funding and Disclosure ...... 29
Figure legends ...... 30
Figures ...... 33
4. Na/K-ATPase Signaling Mediates miR-29b-3p Regulation and Cardiac Fibrosis
Formation in Mice with Chronic Kidney Disease ...... 41
Abstract ...... 43
Introduction ...... 45
Results ...... 46
Discussion ...... 51
Materials and Methods ...... 54
Funding and Disclosure ...... 61
Acknowledgement ...... 62
Tables ...... 63
Figure Legends ...... 65
Figures ...... 69
5. Summary ...... 77
References ...... 79
ix
List of Tables
4. Na/K-ATPase Signaling Mediates miR-29b-3p Regulation and Cardiac Fibrosis
Formation in Mice with Chronic Kidney Disease
Table 1. Echocardiographic data in WT mice ...... 63
Table 2. Echocardiographic data in α1+/- mice ...... 64
x
List of Figures
2. Literature review
Figure1 Structure of Na/K-ATPase...... 5
Figure2 Albers-Post scheme for ion pumping...... 7
3. Characterization of a Long Non-Coding RNA, the Antisense RNA of Na/K-
ATPase Alpha 1 in Human Kidney Cells
Figure 1 Schematic presentation of ATP1A1 and ATP1A1-AS1 gene...... 33
Figure 2 Differential expression and subcellular distribution of ATP1A1-AS1 ...... 34
Figure 3 Epigenetic regulation of ATP1A1-AS1 expression ...... 35
Figure 4 The effect of FOXA1 on ATP1A1-AS1 expression...... 36
Figure 5 Overexpression of ATP1A1-AS1-203 ...... 37
Figure 6 Overexpression of intron 2 of ATP1A1-AS1-203...... 38
Figure 7 Overexpression of ATP1A1-AS1-203 on Src signaling ...... 39
Figure 8 Overexpression of ATP1A1-AS1-203 on cell growth ...... 40
4. Na/K-ATPase Signaling Mediates miR-29b-3p Regulation and Cardiac Fibrosis formation in Mice with Chronic Kidney Disease
Figure 1 PNx-induced miR-29b-3p expression change in WT and α1+/- mice...... 69
Figure 2 Activation of Src and NFκB in mice left ventricle tissue after PNx surgery...... 70
xi Figure 3 Expression and translocation of NFκB in heart tissue after PNx surgery...... 71
Figure 4 The effect of NFκB inhibitor on ouabain-induced miR-29b-3p regulation in
cardiac fibroblasts isolated from WT and α1+/- mice...... 72
Figure 5 Injection of pNaKtide blocks Src activation in mice subjected to PNx surgery. 73
Figure 6 Injection of pNaKtide diminishes PNx-induced decrease in miR-29b-3p
expression ...... 74
Figure 7 Injection of pNaKtide mitigates PNx-induced cardiac fibrosis ...... 75
Figure 8 Injection of pNaKtide attenuates PNx-induced cardiac hypertrophy ...... 76
xii
List of Abbreviations
ATP ...... Adenosine Triphosphate ATP1A1-AS1 ...... ATP1A1 antisense RNA
CKD ...... Chronic Kidney Disease CTS ...... Cardiotonic Steroids
EF ...... Ejection Fraction EGFR ...... Epidermal Growth Factor Receptor ERK...... Extracellular Signal- Regulated Kinase
FS ...... Fractional Shortening
HDAC ...... Histone Deacetylase HF ...... Heart Failure lncRNAs ...... Long Non-Coding RNAs
MBG ...... Marinobufagenin MI ...... Myocardial Infarction
PI3K ...... Phosphatidylinositol-4,5-bisphosphate 3-Kinase PLC ...... Phospholipase C
RAAS ...... Renin-Angiotensin-Aldosterone System RNAi ...... RNA Interference
SAHA ...... Suberoylanilide Hydroxamic Acid
xiii Chapter 1
Introduction
Heart failure is an important public health issue and a leading cause of mortality in the United States (Benjamin, Virani et al. 2018). Tremendous work has been done during the past twenty years to show that Na/K-ATPase and its signaling function plays an important role in regulating cardiac remodeling and cardiac function (Moseley, Cougnon et al. 2004, Tian, Li et al. 2009, Shapiro and Tian 2011, Liu, Bai et al. 2012, Liu, Tian et al. 2016). Our laboratory focuses on studying the effect of Na/K-ATPase on cardiac functions in different animal models and investigating the regulation of Na/K-ATPase expression.
Na/K-ATPase is a transmembrane protein that was discovered in 1957 by Dr. Skou
(Skou 1957). It is a major ion transporter that helps maintain homeostasis of Na+ and K+ concentrations across the cell membrane by hydrolyzing ATP (Skou and Esmann 1992).
Na/K-ATPase ligands, such as cardiotonic steroids (CTS), specifically bind to the extracellular portion of Na/K-ATPase α subunit and cause a conformational change which inhibits the ion transporting activity and ATP hydrolysis (Bagrov, Shapiro et al. 2009). In addition to its canonical ion transporting function, we and other laboratories have shown that α1 Na/K-ATPase can function as a signal transducer. Interaction between CTS and
1 Na/K-ATPase α1 subunit induces activation of Src, PI3K, NFκB, Erk1/2, PLC and other signaling pathways as well as the generation of reactive oxygen species (Kometiani, Li et al. 1998, Liu, Tian et al. 2000, Liu, Mohammadi et al. 2003, Tian, Liu et al. 2003, Liu,
Kesiry et al. 2004, Wang, Haas et al. 2004, Liu, Liang et al. 2005). Activation of these signaling pathways contributes to both physiologic and pathologic changes in the body.
Previous studies have shown the activation of Na/K-ATPase signaling and downstream protein kinase C δ (PKC δ) leads to degradation of friend leukemia integration 1 (Fli-1), which subsequently increases collagen synthesis and stimulate cardiac fibrosis in animals subjected to the 5/6th partial nephrectomy (PNx) (Elkareh, Kennedy et al. 2007, Elkareh,
Periyasamy et al. 2009). Our laboratory has previously shown that Na/K-ATPase α1 related
Src signaling can affect collagen expression through miR-29b-3p dysregulation. The current study was designed to test the in vivo effect of Na/K-ATPase signaling in regulating the expression of miR-29b-3p and tissue fibrosis in a mouse model of CKD. We will also investigate the mechanism of miR-29b-3p regulation by Na/K-ATPase α1 related signaling.
In addition to the signaling function listed above, appropriate Na/K-ATPase expression is also important for maintaining normal cell and body function. Dysregulation of Na/K-ATPase expression has been observed in different diseases. In the failing heart, both Na/K-ATPase expression (Norgaard, Bagger et al. 1988) and pumping activity
(Schwinger, Wang et al. 1999) were found decreased and its expression level was correlated with the patients ejection fraction (Norgaard, Bagger et al. 1988, Ishino, Botker et al. 1999). Na/K-ATPase reduction were also documented in patients or animal models with CKD (Drummond, Sayed et al. 2014), aging (Fraser and Arieff 2001), diabetes with
2 hypertension (Chen, Yuan et al. 1993, Straub, Hall et al. 1996), neurological disorders
(Kawamoto, Munhoz et al. 2008), and cancer (Litan and Langhans 2015).
Previous studies have discovered a panel of transcription factors, hormones, growth factors, and lipids that can regulate Na/K-ATPase expression (Li and Langhans 2015). Our recent work identified a natural antisense gene (ATP1A1-AS) as a novel endogenous regulator of Na/K-ATPase expression.
The ATP1A1 antisense RNA (ATP1A1-AS1) is one type of long non-coding
RNAs, which is located on the opposite strand of the sense ATP1A1 gene and was first reported in a human genomic study (Strausberg, Feingold et al. 2002). My dissertation is aimed to characterize this ATP1A1-AS1 gene, and its role in regulating the Na/K-ATPase
α1 expression and its signaling function. My dissertation also includes a study that investigates how Na/K-ATPase reduction regulates Na/K-ATPase signaling and cardiac function in a CKD animal model.
3 Chapter 2
Literature review
2.1 Na/K-ATPase biology
2.1.1 Structure of Na/K-ATPase
Na/K-ATPase is a transmembrane protein that was discovered in 1957 by Dr. Skou that functions as an ion pump (Skou 1957). By hydrolyzing ATP, it transports 3 sodium ions out of the cell and transports 2 potassium ions into the intracellular space. This process helps maintaining the cellular resting potential and regulates cellular volume. The essential components of the Na/K-ATPase protein are the α subunit and β subunit (Figure 1). In some tissues, the α-β heterodimer can be associated with a γ subunit, which belongs to the
FXYD family (Tokhtaeva, Clifford et al. 2012).
The α subunit of Na/K-ATPase has 10 transmembrane segments (TM1-TM10) and three intracellular domains (Skou and Esmann 1992). The extracellular segments of the α subunit form a binding site for CTS, the ligands of Na/K-ATPase. The intracellular portion of Na/K-ATPase forms three major domains (N-domain, P-domain, and A-domain). The
Intracellular loop that links the TM4–TM5 forms the nuclear binding domain (N-domain) and phosphorylation domain (P-domain). N-domain has a “pocket” structure that is considered as the binding site for ATP. The P-domain is located on the proximal and distal
4 parts of intracellular loop. A phosphate molecule from ATP is transiently transferred on the aspartyl residue 376 of the DKTGT motif of P-domain during ATP hydrolysis. The actuator domain (A-domain) is constituted by the cytoplasmic NH2-terminal and TM2–
TM3 intracellular loop (Sweadner and Donnet 2001, Kanai, Ogawa et al. 2013).
Figure 1. Structure of Na/K-ATPase. The γ subunit is tissue specific and belongs to the FXYD family. The FXYD2 is highly conserved among different species.
The major role of α subunit is hydrolyzing ATP and transporting the cations across cell membrane. The function of β subunit is not well understood, but it is essential for transporting the α subunit onto the plasma membrane (Geering 1991, Hilbers, Kopec et al.
2016). The FXYD subunit is not an integral part of the Na/K-ATPase, but is associated with specific domains of the αβ subunit complex and helps to modulate the catalytic properties of the Na/K-ATPase (Crambert, Fuzesi et al. 2002).
There are four α (α1, α2, α3, and α4) isoforms and three β (β1, β2, and β3) isoforms and thus allowing different combinations of αβ complexes in different tissues. The α1 isoform is expressed ubiquitously in all cells, α2 and α3 are predominantly found in
5 myocytes and neurons, respectively; and α4 is testis-specific. The expression of β1 is also ubiquitous. The β2 and β3 isoforms are expressed in brain, cartilage, and erythrocytes, whereas β2 can also be found in cardiac tissues and β3 can be observed in lung tissues.
These different isoforms also alter the Na/K-ATPase sensitivity to its ligands, CTS, and result in different activity in both pumping and enzyme function (Bagrov, Shapiro et al.
2009).
2.1.2 Ion pumping function of Na/K-ATPase
The Na/K-ATPase is a member of the P-type ATPase family of membrane- incorporated proteins. Conformational changes of the Na/K-ATPase α subunit was discovered in the ion transporting activity. As shown in Figure 2, the α subunit, in the presence of Na+ and Mg2+, is first phosphorylated by ATP. This is followed by the occlusion of three cytosolic Na+ ions. This high-energy E1P form of the enzyme, loaded with Na+ ions, undergoes a conformational change to the low-energy E2P form. When the sodium cation sites are exposed to the extracellular medium, Na+ ions are released, and in the presence of potassium ions, the E2P form is dephosphorylated. Dephosphorylation is followed by occlusion of two K+ ions to specific potassium binding sites. Uptake of K+ ions leads to the transition of the E2 form to the E1 form, which is accelerated by ATP.
This transition is followed by the release of K+ ions to the intracellular medium. Then the
E1 form with bound ATP undergoes the cycle again. CTS can specifically bind to the extracellular portion of the Na/K-ATPase subunit at the E2P conformation, which blocks the enzyme from entering the next conformation, and therefore inhibits its ion transporting activity (Lingrel 1992).
6
Figure 2. Albers-Post scheme for ion pumping.
2.1.3 Ligands of Na/K-ATPase
Digitalis compounds such as digoxin have been used in treatment of congestive heart failure for more than 200 years due to their positive inotropic effect (Grupp, Im et al.
1985, Smith 1988). It was until 1960s, these digitalis compounds were found to be specific ligands of Na/K-ATPase (Braunwald and Klocke 1965). Since the structurally similar substances were also found in humans and animals, they are also termed as cardiotonic steroids (CTS). Binding of CTS to the Na/K-ATPase in the cardiac myocytes causes inhibition of Na/K-ATPase activity and result in increased intracellular Na+ and Ca2+ that leads to a positive inotropic effect (Smith 1988).
CTS are a group of chemical compounds sharing a similar core structure of either five or six lactone and can be classified into cardenoildes (Buckalew 2015) or bufadenolides (Steyn and van Heerden 1998), respectively . In cardenolides CTS, a five- membered lactone ring is connected to a steroidal nucleus at position 17 and a hydroxyl
7 group is at position 14. On the other hand, bufadienolides CTS have a six-membered lactone ring in position 17 and some also have a 14-15 epoxide.
2.1.4 Signaling function of Na/K-ATPase
In addition to the canonic pumping function, ample evidence have suggested that
Na/K-ATPase may function as a signaling transducer (Xie 2003, Bagrov, Shapiro et al.
2009, Liu, Lilly et al. 2018). Experiments from Dr. Xie’s laboratory found that there is a
“pool” of Na/K-ATPases that reside in the caveolae structure and directly associate with different signaling molecules (Wang, Haas et al. 2004, Liang, Tian et al. 2007). Exposure of Na/K-ATPase to CTS such as ouabain or MBG induces rapid phosphorylation of Src and EGFR (Haas, Askari et al. 2000, Haas, Wang et al. 2002). In addition to the EGFR, other signaling proteins seem to be recruited including phospholipase C, TRP proteins,
PI(3)K, and several isoforms of PKC (Kometiani, Li et al. 1998, Liu, Tian et al. 2000, Liu,
Mohammadi et al. 2003, Tian, Liu et al. 2003, Liu, Kesiry et al. 2004, Wang, Haas et al.
2004, Liu, Liang et al. 2005). Further studies showed that Na/K-ATPase binds with Src and maintains Src in an inactive form. The binding of CTS to the Na/K-ATPase induces a conformation change that, in turn, alters the interaction between the Na/K-ATPase and Src and allows Src to become activated. This active Src is then able to phosphorylate other downstream signaling proteins (Tian, Cai et al. 2006). GST pull-down assays (Tian, Cai et al. 2006) showed that there are at least two contacting sites between α1 Na/K-ATPase and Src: CD2 domain of α1 with Src SH2 domain, and the CD3 domain of α1 with Src kinase domain. Further mapping of this interaction has identified the 20 amino acid
NaKtide sequence in the nucleotide-binding domain of α1 Na/K-ATPase being responsible
8 for the direct interaction between α1 CD3 and Src kinase domain (Tian, Cai et al. 2006, Li,
Cai et al. 2009). The synthesized NaKtide can interact with and inhibits Src.
Na/K-ATPase also directly regulate PI3K/Akt/mTOR pathway, which could be Src independent (Liu, Li et al. 2006, Liu, Zhao et al. 2007, Wu, Li et al. 2015). CTS treatment activates PI3Kα but not PI3Kγ in cultured myocytes and causes hypertrophic growth (Liu,
Zhao et al. 2007). These studies also found that low concentration of ouabain (< 40 nM) blocks Na/K-ATPase α2 related signaling blocks transverse aortic constriction (TAC)- induced pathological hypertrophy in mice. However, higher concentration of ouabain activates Na/K-ATPase α1 related signaling that leads to pathological hypertrophy and cell death (Wu, Li et al. 2015).
2.2 Heart failure and cardiac remodeling
Heart failure (HF) is associated with significant high morbidity and mortality that largely attributable to cardiac remodeling. Cardiac remodeling is defined as manifest changes in size, mass, geometry and function of the heart (Cohn, Ferrari et al. 2000). The term "remodeling" was used for the first time in 1982 by Hockman and Buckey, in a myocardial infarction (MI) model (Hochman and Bulkley 1982). It was aimed to characterize the replacement of infarcted tissue with scar tissue (Hochman and Bulkley
1982) . The term was then used in scientific articles on morphological changes following acute MI, particularly progressive increase of the left ventricular cavity (Pfeffer, Pfeffer et al. 1985, Pfeffer and Braunwald 1990).
Cardiac remodeling can be classified into physiologic or pathologic. Physiologic remodeling, which is defined as compensatory changes that improves heart function.
Pathologic remodeling refers to the pathologic conditions caused by MI, uremic
9 cardiomyopathy, aortic stenosis, hypertension, inflammatory heart muscle disease
(myocarditis), or idiopathic dilated cardiomyopathy, in which the heart function was deteriorated due to the cardiac remodeling (Xie, Burchfield et al. 2013, Azevedo, Polegato et al. 2016).
A number of cellular changes are involved in cardiac remodeling including myocyte hypertrophy (Grossman, Jones et al. 1975, Anversa, Olivetti et al. 1991), loss of myocytes due to apoptosis (Sharov, Sabbah et al. 1996, Teiger, Than et al. 1996, Olivetti,
Abbi et al. 1997) or necrosis (Tan, Jalil et al. 1991), fibroblast proliferation (Villarreal,
Kim et al. 1993), and fibrosis (Anderson, Sutton et al. 1979, Weber, Pick et al. 1990). In addition, abnormalities of the microvasculature, such as reductions in capillary density, thickening of the walls of the arterioles, and inadequate angiogenesis are also involved in the pathogenesis of cardiac remodeling (Gerdes, Callas et al. 1979, Rakusan, Flanagan et al. 1992).
As part of a compensatory process to maintain stroke volume, surviving myocytes become elongated or hypertrophied following an injury to the heart, especially after loss of cardiomyocyte number (Grossman, Jones et al. 1975, Anversa, Olivetti et al. 1991). In cardiac myocytes, stretching of cell membranes resulting from cardiac overload will induce the expression of hypertrophy associated genes and lead to synthesis of new contractile proteins and the assembly of new sarcomeres (Francis and McDonald 1992).
Cardiac fibrosis can be categorized into two types: reactive fibrosis and replacement fibrosis (also called reparative fibrosis) (Weber and Brilla 1992). Replacement fibrosis often occurs after MI when large numbers of cardiac myocytes undergo necrosis.
Myocyte necrosis triggers a series of events including immune cell infiltration,
10 inflammation, new vessel formation, removal of necrotic tissue, and eventually the replacement of damaged tissue with collagen-dominated fibrotic tissue that prevents cardiac muscle from rupture (Prabhu and Frangogiannis 2016). In addition to scar formation at the infarcted area, the remote non-infarcted regions can develop fibrosis in the interstitial spaces, which is referred to as reactive fibrosis or interstitial fibrosis (Weber and
Brilla 1992, Cleutjens, Verluyten et al. 1995, Sun and Weber 2000, Krenning, Zeisberg et al. 2010). Interstitial fibrosis also occurs in other disease conditions related with activation of renin-angiotensin-aldosterone system (RAAS) (Brilla 2000, Schnee and Hsueh 2000,
Zhang, Zhang et al. 2010), endothelin-1 (Shi-Wen, Rodriguez-Pascual et al. 2006, Adiarto,
Heiden et al. 2012, Wang, Guo et al. 2015), TGF-β (Leask and Abraham 2004, Bujak and
Frangogiannis 2007, Bujak, Ren et al. 2007), TNF- (Zhang, Chancey et al. 2011,
Duerrschmid, Crawford et al. 2013), NFB (Chen and Greene 2004, Gordon, Shaw et al.
2011), and other profibrotic signaling pathways (Biernacka and Frangogiannis 2011, Tao,
Yang et al. 2016).
2.3 Na/K-ATPase and cardiac remodeling
Na/K-ATPase is an important protein in cardiac tissue that is critical to maintain the cellular ion homeostasis. Experimental data showed that over activation of Na/K-
ATPase signaling by increased CTS levels in disease conditions could stimulate myocytes hypertrophic growth as well as cardiac fibrosis (Elkareh, Kennedy et al. 2007, Wansapura,
Lasko et al. 2011) . It was also found that the amount of Na/K-ATPase in the heart tissue affect the ion pumping (Schwinger, Wang et al. 1999, Liang, Cai et al. 2006) function as well as its signaling function (Liu, Bai et al. 2012, Drummond, Sayed et al. 2014,
11 Drummond, Fan et al. 2018). These changes in turn contribute to the pathological change of the heart.
The expression pattern of the three Na/K-ATPase α isoforms depends on species and development stage. In rat, αl is present at all development stages, while α3 is present during fetal life and is replaced by α2 during adult life (Orlowski and Lingrel 1988, Shyjan and Levenson 1989). In humans, the α1 and α3 are predominantly expressed in fetal heart, and all three isoforms are expressed at comparable level in adult heart (Gilmore-Hebert,
Schneider et al. 1989, Lucchesi and Sweadner 1991, Shamraj, Melvin et al. 1991).
Early studies found that Na/K-ATPase content was significantly reduced in heart tissue from patients with dilated cardiomyopathy and heart failure (Kjeldsen, Bjerregaard et al. 1988, Norgaard, Bagger et al. 1988, Schwinger, Bohm et al. 1990, Schwinger, Bohm et al. 1992, Shamraj, Grupp et al. 1993). In addition to the protein amount, the ATPase activity of Na/K-was also found decreased in homogenates and membranes from failing human hearts (Schwinger, Wang et al. 1999). Functional studies showed that the decreased
Na/K-ATPase is related with the decreased left ventricular ejection fraction (Norgaard,
Bagger et al. 1988, Ishino, Botker et al. 1999). Later studies in animal models showed that the reduction of Na/K-ATPase expression occurs early in the development of heart failure, probably before heart failure symptoms are present (Hanf, Drubaix et al. 1988, Zahler,
Gilmore-Hebert et al. 1996, Zobel, Brixius et al. 1998), indicating that reduction of Na/K-
ATPase in the heart may contribute to heart failure development. Our recent publications have shown that reduction of Na/K-ATPase attenuated Src and mTOR pathways in animals subjected to CTS infusion or partial nephrectomy (PNx) surgery and caused significant cardiac cell apoptosis (Liu, Bai et al. 2012, Drummond, Sayed et al. 2014).
12 The elevation of circulating levels of these endogenous CTS was reported in heart failure patients and was correlated with the severity of heart dysfunction (Gottlieb,
Rogowski et al. 1992, Bagrov, Fedorova et al. 1995, Gonick, Ding et al. 1998, Simonini,
Pozzoli et al. 2015). Other diseases such as renal artery stenosis (Tian, Haller et al. 2010), preeclampsia (Lopatin, Ailamazian et al. 1999), myocardial ischemia/infarction (Bagrov,
Fedorova et al. 1998), and diabetes mellitus (Straub, Hall et al. 1996) were also observed in association with elevated levels of endogenous compounds in human plasma samples.
Mechanistically, in animals or cultured cells with regular Na/K-ATPase content, increased CTS can stimulate cell proliferation and protect cells from death (Abramowitz,
Dai et al. 2003, Li, Zelenin et al. 2006, Tian, Li et al. 2009) by activating Na/K-ATPase associated Src, PI3K and NFkB signaling pathway. Prolonged activation of these signaling pathways in certain disease models was demonstrated to cause cardiac remodeling such as hypertrophy (Wansapura, Lasko et al. 2011) and fibrosis (Elkareh, Kennedy et al. 2007).
However, in animals with reduced Na/K-ATPase, elevated endogenous CTS activates caspase 9 and 3 which induces myocyte apoptosis, ventricular dilation, and decreased cardiac function (Liu, Bai et al. 2012).
Activation of Na/K-ATPase signaling also contributes to the formation of cardiac fibrosis in animals subjected to CTS infusion or PNx surgery through activation of Src and
PKCδ. Activation of Src stimulates the fibroblast proliferation and activation of PKCδ induces phosphorylation of Friend leukemia integration-1 (Fli-1) (Elkareh, Kennedy et al.
2007). Fli-1 is a transcription factor that negatively regulates collagen mRNA synthesis, while phosphorylation of Fli-1 can cause degradation of Fli-1 and subsequently induces collagen synthesis (Elkareh, Periyasamy et al. 2009). Interestingly, TGF-β and its
13 downstream component Smad 2/3 or Smad 4 was not activated in the heart tissue of these animals (Elkareh, Kennedy et al. 2007). In our laboratory, we showed that CTS, such as ouabain and MBG, decrease miR-29b-3p, an anti-fibrotic microRNA, and lead to an increase in collagen synthesis through a Src-related signaling pathway in cardiac fibroblasts (Drummond, Hill et al. 2015). In chapter 4 of this dissertation, we extended the mechanistic study of miR-29b-3p expression following Na/K-ATPase signaling activation.
We appreciated the importance of NFkB activation in miR-29b-3p regulation (Drummond,
Fan et al. 2018).
2.4 Regulation of Na/K-ATPase
Given the importance of Na/K-ATPase in ion pumping and cell signal transduction, the regulation of Na/K-ATPase expression is essential for normal cell function. Studies from the last century have shown that the spatial and temporal expression of Na/K-ATPase is partially regulated at the transcriptional level. Numerous transcription factors, hormones, growth factors, lipids, and extracellular stimuli has been characterized and were reviewed by (Li and Langhans 2015). Each subunit and isoform may have different responses to these stimulators. The responses are also organ, cell type, and species dependent. For example, dexamethasone can increase α2, but has no effect on α1, α3, and β in cultured neonatal rat cardiac myocytes (Orlowski and Lingrel 1990). However in in rat capsule- epithelium of lenses, dexamethasone will decrease α1 expression (Xie and Askari 2002,
Xie, Yan et al. 2010).
Recent advances in genome-wide sequencing have revealed that the majority of the human genome was transcribed, but only a small portion of the transcribed RNAs actually codes proteins (Djebali, Davis et al. 2012, Hangauer, Vaughn et al. 2013, de Hoon, Shin et
14 al. 2015). The RNAs lacking protein coding information were defined as non-coding
RNAs, which, depending on their length, were further divided into short non-coding RNAs
(less than 200 nucleotides) and long non-coding RNAs (lncRNAs) (Ponting, Oliver et al.
2009, Ulitsky and Bartel 2013, Dykes and Emanueli 2017). It was estimated that lncRNAs only make up 0.03-0.2% of total RNA by mass (Palazzo and Lee 2015). Many of the discovered lncRNAs are located on the antisense strand of well-defined transcriptional units, which are usually called natural antisense RNA (Papaioannou, Nicolet et al. 2017).
Some identified natural antisense genes were found to regulate their sense protein-coding gene expression (Wahlestedt 2013) or act in cellular processes such as cell-cell communication (Geisler and Coller 2013, Villegas and Zaphiropoulos 2015). The ATP1A1 antisense RNA (ATP1A1-AS1) is a natural antisense gene of Na/K-ATPase α1 (ATP1A1).
This antisense gene is located on the opposite strand of the sense ATP1A1 gene and was first reported in a human genomic study (Strausberg, Feingold et al. 2002). In this dissertation, I investigate the effect ATP1A1-AS1 on expression of Na/K-ATPase α1
(ATP1A1) and present the results in chapter 3.
15 Chapter 3
Characterization of A Long Non-Coding RNA, The Antisense RNA of Na/K-ATPase Alpha 1 In Human Kidney Cells
Xiaoming Fan1, Usman M. Ashraf2, Christopher A. Drummond1,3, Huilin
Shi1, Xiaolu Zhang1, Sivarajan Kumarasamy2, and Jiang Tian1, *
1 Department of Medicine at the University of Toledo, Toledo, OH 43614, United
States of America
2 Department of Physiology and Pharmacology, Center for Hypertension and
Personalized Medicine, at the University of Toledo, Toledo, OH 43614, United States of
America
3 MPI Research, Mattawan, MI 49071, United States of America
* Correspondence: [email protected]; Tel.: (419) 383-3510
This work has been published in International Journal of Molecular Sciences 19(7):
2123.
16 Abstract
Non-coding RNAs have been suggested to be important regulators of protein- coding genes. The objective of this study is to investigate the function of a long non-coding antisense RNA, ATP1A1-AS1, in human kidney cells. We have characterized an antisense long non-coding RNA (ATP1A1-AS1) that is located on the opposite strand of the sense gene of the Na/K-ATPase alpha1, which has at least 4 splice variants expressed in human embryonic kidney cells (HEK293 cells) as well as in adult kidney cells (HK2 cells).
Expression of the ATP1A1-AS1 can be regulated by histone acetylation as well as DNA methylation. Overexpression vector of ATP1A1-AS1 exons or introns was constructed by cloning their cDNA sequence into a pcDNA3.1(-B) plasmid and was transfected into human kidney cells to test their physiological role. The result showed that overexpression of the ATP1A1-AS1 transcript in HK2 cells reduced the expression of Na/K-ATPase α1
(ATP1A1) gene by about 20% (p<0.05). It also reduced the Na/K-ATPase α1 protein level by about 22% (p<0.05). Overexpression of this antisense RNA transcript attenuated ouabain-induced Src activation, and subsequently inhibited cell proliferation and potentiated ouabain-induced cell death. These results demonstrate that ATP1A1-AS1 gene can serve as a moderate negative regulator of Na/K-ATPase and modulate the Na/K-
ATPase-related signaling pathways in human kidney cells.
Keywords: Long non-coding RNA; ATP1A1-AS1; Na/K-ATPase; Src, Signaling transduction.
17 Introduction
Recent advances in genome-wide sequencing has revealed that majority of the human genome was transcribed, but only a small portion of the transcribed RNAs actually codes proteins (Djebali, Davis et al. 2012, Hangauer, Vaughn et al. 2013, de Hoon, Shin et al. 2015). The RNAs lacking protein coding information were defined as non-coding
RNAs, which, depending on their length, were further divided into short non-coding RNAs
(less than 200 nucleotides) and long non-coding RNAs (lncRNAs) (Ponting, Oliver et al.
2009, Ulitsky and Bartel 2013, Dykes and Emanueli 2017). lncRNAs have some features similar to protein coding messenger RNAs (mRNAs), such as the structure of introns and exons. Many lncRNAs have polyA tail and express splice variants (Guttman, Amit et al.
2009, Hangauer, Vaughn et al. 2013). However, the expression level of lncRNAs is much lower comparing to mRNA, rRNA, or tRNA. It was estimated that lncRNAs only makes up 0.03-0.2% of total RNA by mass (Palazzo and Lee 2015). Many of the discovered lncRNAs are located on the antisense strand of well-defined transcriptional units, which are usually called natural antisense RNA (Papaioannou, Nicolet et al. 2017). Some identified natural antisense genes were found to regulate their sense protein-coding gene expression (Wahlestedt 2013) or act in cellular process such as cell-cell communication
(Geisler and Coller 2013, Villegas and Zaphiropoulos 2015). However, the physiological function of most lncRNAs or natural antisense RNAs have not been elucidated.
The ATP1A1 antisense RNA (ATP1A1-AS1) is a natural antisense gene of Na/K-
ATPase α1 (ATP1A1). This antisense gene is located on the opposite strand of the sense
ATP1A1 gene and was first reported in a human genomic study (Strausberg, Feingold et al. 2002). The ATP1A1-AS1 gene has at least 4 splice variants (ATP1A1-AS1-201,
18 ATP1A1-AS1-202, ATP1A1-AS1-203, and ATP1A1-AS1-204). ATP1A1-AS1-203 and
ATP1A1-AS1-204 are partially overlapping with the sense ATP1A1 gene. At least 27 human tissues including kidney, heart, and blood were demonstrated to express the
ATP1A1-AS RNA transcripts (Fagerberg, Hallstrom et al. 2014). ATP1A1-AS1 expression is higher in kidney, thyroid, and intestine (Fagerberg, Hallstrom et al. 2014,
Duff, Olson et al. 2015).
Na/K-ATPase is a transmembrane protein that was discovered in 1957 by Dr. Skou
(Skou 1957). In addition to its canonical ion transporting function, the Na/K-ATPase α1 was found to be associated with other signaling proteins and function as a signal transducer
(Xie and Askari 2002, Xie 2003, Aperia, Akkuratov et al. 2016). Na/K-ATPase function is closely related with cardiac function and has been recognized as a potential treatment target for heart failure (Rathore, Curtis et al. 2003, Chan, Lazarus et al. 2010). Data from clinical research showed that the cardiac contractility is positively correlated with Na/K-ATPase levels in heart failure patients (Norgaard, Bagger et al. 1988, Ishino, Botker et al. 1999).
Experimental animal models also demonstrated that reduction of Na/K-ATPase α1 potentiates cardiac cell apoptosis and worsens cardiac function (Moseley, Cougnon et al.
2004, Yan, Haller et al. 2012, Drummond, Sayed et al. 2014). Na/K-ATPase expression can be regulated by multiple transcriptional factors and a variety of chemical compounds
(Li and Langhans 2015). The current work is aiming to characterize this ATP1A1-AS1
RNA and its role in regulation of Na/K-ATPase α1 expression and its signaling function in human kidney cells.
19 Results
Differential expression and subcellular distribution of ATP1A1-AS1 splice variants in human kidney cells
The ATP1A1-AS1 gene locates in the region of 116,392,247-116,418,622 on the reverse strand of human chromosome 1 (Figure 1) based on Ensembl GRCh38.p12
(http://www.ensembl.org). To assess the expression level of each splice variants, we synthesized specific primers corresponding to the 4 known transcripts of ATP1A1-AS1 as described in Material and Method section. As shown in Figure 2A, all 4 splice variants can be detected in HK2 cell lysates, while the ATP1A1-AS1-203 expression is relatively higher than the other three transcripts. A similar expression pattern was also observed in HEK293 cells, another human kidney cell line (Figure 2B). However, the overall expression level of the antisense transcripts are much lower compared to the sense gene, ATP1A1. We also examined the subcellular distribution of these antisense transcripts in HK2 cells and found that these transcripts were present in both cytosol and nuclear fraction (Figure 2C).
Epigenetic regulation of ATP1A1-AS1 expression
To understand the regulation mechanism of ATP1A1-AS1 expression, we treated
HK2 cells with a histone deacetylase (HDAC) inhibitor, suberoylanilide hydroxamic acid
(SAHA), at concentrations of 10 nM, 100 nM, 1 M, and 10 M for 48 h. We then measured the ATP1A1-AS1-203 levels as the representative of ATP1A1-AS1 in the above drug-treated cell lysates. As shown in Figure 3A, SAHA treatment at lower concentrations
(10 and 100 nM) had no significant effect on the expression of the sense or antisense gene expression. However, SAHA at higher concentrations (1 and 10 M) significantly increased the expression of ATP1A1-AS1-203. It also increased the expression of
20 ATP1A1, but to a much lesser degree. We also treated the cells with a DNA methylation inhibitor, Decitabine, at concentration of 5 nM, 50 nM, 500 nM, and 5 M for 48 h to test if DNA methylation regulates the ATP1A1-AS1 expression. The result showed that inhibition of DNA methylation had significant effect on upregulating the antisense gene
(ATP1A1-AS1) expression, but it did not affect the ATP1A1 gene expression (Figure 3B).
These results suggest a differential effect of methylation and acetylation on the regulation of the sense and antisense gene.
We also identified a FOXA1 binding site on the upstream of ATP1A1-AS1 gene based on the sequence information. To test if FOXA1 regulates the antisense gene expression, we cloned FOXA1 cDNA into the pcDNA3.1(-B) vector and transfected into
HK2 cells. As shown in Figure 4, overexpression of FOXA1 induced only a slight increase in the expression of ATP1A1-AS1-203 and a slight decrease in the expression of ATP1A1 in these cells, which had no statistical significance.
ATP1A1-AS1 regulates the sense Na/K-ATPase α1 gene expression and protein synthesis in HK2 cells
To test the role of ATP1A1-AS1 on the regulation of its sense gene, we constructed a plasmid vector that overexpresses ATP1A1-AS1-203, which is the highest RNA transcript of the ATP1A1-AS1 gene, and transient transfected into the cultured HK2 cells for 48 h and measured the Na/K-ATPase α1 mRNA and protein levels using RT-qPCR and
Western blot, respectively. As shown in Figure 5, overexpression of ATP1A1-AS1-203 caused about 20% decrease (p<0.05) in ATP1A1 mRNA levels and a 22% decrease in
Na/K-ATPase α1 protein levels in these cells, indicating that the antisense gene is a negative regulator of Na/K-ATPase α1.
21 In addition to the exons of ATP1A1-AS1-203 which has complementary sequence to the mRNA of Na/K-ATPase α1, we also found that the intron 2 of this transcript contains sequence complementary to the sense gene. Since the introns of lncRNAs may also involve in regulation of gene expression, we examined the presence of the intron 2 of RNA transcript and its role in regulating the sense gene expression in HK2 cells. As shown in
Figure 6A, the intron 2 of ATP1A1-AS1-203 was detectable in the total RNA extracted from HK2 cell lysates. However, overexpression of this intron 2 via 48 h pcDNA3.1(-B) plasmid transfection in HK2 cells had only slight effect on the expression of sense ATP1A1 gene (Figure 6B).
Overexpression of ATP1A1-AS1 regulates Na/K-ATPase-related signaling and cell proliferation.
Our previous studies (Liang, Cai et al. 2006, Tian, Li et al. 2009, Li and Chen 2013) have shown that reduction of Na/K ATPase α1 attenuates cardiotonic steroids (CTS)- induced Src activation and potentiates CTS-induced cell growth inhibition. To study the effect of ATP1A1-AS1 on the Na/K-ATPase-related signaling function, we transient transfected HK2 cells with the ATP1A1-AS1-203 overexpression vector for 24h followed by ouabain treatment for 15 min. Empty vector transfected cells were used as control. The cell lysate was then collected to probe for Src phosphorylation at tyrosine 418 (pSrc418), an indicator of Src activation. As shown in Figure 7, ouabain treatment at 50 nM induced significant increase in Src phosphorylation in cells transfected with empty vector, whereas in the cells that were overexpressed ATP1A1-AS1-203, ouabain failed to induce Src activation.
22 To examine the role of ATP1A1-AS1 on cell proliferation, we transfected the cells with ATP1A1-AS1-203 overexpression vector. The cells were then incubated in an
Incucyte incubator equipped with a camera to monitor the cell growth for 96 h. To investigate the cell growth in the presence of ouabain, HK2 cells with 48 h ATP1A1-AS1-
203 overexpression were treated with 50 nM ouabain for another 48 h. Cell growth was also monitored for a total of 96 h. As shown in Figure 8, overexpression of ATP1A1-AS1-
203 inhibited cell proliferation in the absence or presence of ouabain. In some areas, we also observed that addition of ouabain induced cell death in the ATP1A1-AS1-203 overexpressed cells. These results are in line with our previous observations in the setting of decreased Na/K-ATPase α1 expression (Tian, Li et al. 2009, Yan, Haller et al. 2012).
Discussion
In this report, we examined the splice variants and subcellular distribution of a newly identified antisense lncRNA, ATP1A1-AS1. Our result, for the first time, showed that ATP1A1-AS1 is a potential negative regulator of its sense gene, Na/K-ATPase α1, in human kidney cells. It is important to note that decrease of Na/K-ATPase is a common phenomenon in patients with congestive heart failure (Norgaard, Bagger et al. 1988, Semb,
Lunde et al. 1998), aging (El-Mallakh, Barrett et al. 1993, Poehlman 1993, Maurya and
Prakash 2013), diabetes with hypertension (Clerico and Giampietro 1990, Chen, Yuan et al. 1993, Tirupattur, Ram et al. 1993), and neurological disorders (Harik, Mitchell et al.
1989, Liguri, Taddei et al. 1990). Reduction of cardiac Na/K-ATPase was related with a decrease in cardiac contractile function in humans and in animal models (Norgaard, Bagger et al. 1988, Ishino, Botker et al. 1999, Moseley, Cougnon et al. 2004). Consistently, the current work showed that increased expression of ATP1A1-AS1 gene attenuated the
23 ouabain-induced Src activation and inhibited cell growth or potentiate ouabain-induced cell death.
However, we noted that the change in Na/K-ATPase α1 expression by overexpression of antisense ATP1A1-AS1 transcript is moderate in these cells. Our data do show that increased ATP1A1-AS1 expression can alter the Na/K-ATPase-related signaling function and cell proliferation but considering the much lower expression level of the antisense gene relative to the sense gene in normal conditions, the physiological role of ATP1A1-AS1 remains to be elucidated. It is also not clear whether Na/K-ATPase α1 is a specific and the only target of ATP1A1-AS1. On the other hand, molecular mechanisms for antisense RNA-induced sense gene regulation are not fully understood. Mechanisms such as RNA interference (RNAi) by formation of double strand RNA or steric clashes induced by antisense RNA may exist to regulate the corresponding sense gene (Munroe and Zhu 2006, Faghihi and Wahlestedt 2009, Li and Ramchandran 2010). More recently, investigators hypothesized that the antisense RNA could control the quality and quantity of the sense gene by producing endogenous siRNAs (Wight M 2013). In addition, our data showed that the alternative splicing of antisense gene exist in human kidney cells, which may also play a role in regulating the sense gene expression. Identifying these mechanisms in the future will provide more effective tools to manipulate the Na/K-ATPase α1 expression and protect normal cardiac function in humans.
Previous studies have shown that chromatin state such as DNA methylation and histone acetylation modification are important regulators of lncRNAs expression. In the current study, we observed that histone acetylation and DNA methylation had differential effects on ATP1A1-AS1 and ATP1A1 expression. The ATP1A1-AS1 gene expression was
24 more responsive to the epigenetic modification, especially to the change in DNA methylation, whereas the effect of DNA methylation on ATP1A1 gene expression was modest. This observation is consistent with previous findings that DNA methylation change did not affect the ATP1A1 expression (Henriksen, Kjaer-Sorensen et al. 2013,
Selvakumar, Owens et al. 2014). However, based on the data released from
ENCODE/HAIB study (https://www.ncbi.nlm.nih.gov/geo/info/ENCODE.html), in human kidney cells, there are 12 DNA methylation sites with 7 methylated, 1 partially methylated, and 4 unmethylated on the ATP1A1 coding strand, while on the ATP1A1-AS1 coding strand, there are 10 DNA methylation sites with 5 methylated, 1 partially methylated and 4 unmethylated. Therefore, the number of methylation sites or methylated nucleotide number alone may not explain the differential regulation effect on this pair of sense/antisense gene. In addition, even though the sequence of ATP1A1-AS1 data indicate a transcription factor binding site for FOXA1, our experimental data showed that overexpression of FOXA1 failed to regulate the ATP1A1-AS1 expression.
In summary, the current findings showed that the ATP1A1-AS1 can negatively regulate its sense gene expression and affect the Na/K-ATPase signaling function in human kidney cells. However, it merits further studies to fully understand the physiological role of this antisense RNA.
Materials and Methods
Cell culture
Human kidney cells (HK2 cell line and HEK293 cell line) were purchased from
American Type Culture Collection (ATCC) and cultured in Dulbecco's modified Eagle's
25 medium (HK2 cells) or Eagle’s minimum essential medium (HEK293 cells) supplemented with 10% fetal bovine serum (FBS), 100 units/ml penicillin, and 100 μg/ml streptomycin in a humidified incubator with 5% CO2.
Quantitation of ATP1A1-AS1 expression using RT-qPCR
Total RNA was extracted from cultured HK2 or HEK293 cells using RNeasy Mini
Kit from Qiagen (Cat. No.: 74104) following the protocol provided by the manufacturer.
About 1 g of extracted RNA was used for cDNA synthesis with the RT2 First Strand cDNA synthesis kit from Qiagen (Cat. No.: 330404). Expression of ATP1A1-AS1 were quantified by RT-qPCR. The primers used for RT-qPCR were as following: ATP1A1-AS1-
201 (Forward: TTTGCGCTAACGATGAGAAC; reverse:
GCATTTCCAGATGCATGGT), ATP1A1-AS1-202 (Forward:
GGGCTGAGAGCTAAGGAGTG; reverse: ATGGGCATTTCCTCCTGAT), ATP1A1-
AS1-203 (Forward: AGCGGTCATCCCAGTCCAC; reverse:
CCAGTGTGTGTCCCAATCCC), and ATP1A1-AS1-204 (Forward:
GTTCTCAGCCAGAATCACAAACTT; reverse:
GATGAGAGAAAGATACGCCAAAAT), Intron 2 of ATP1A1-AS1-203 (forward:
GTCTCTGAAATCAACCTCAACC, reverse: ACTAAATTCCTTCTCCCCACC).
GAPDH was used as internal control. Primer pair for GAPDH was from Qiagen (Cat No.:
PPH00150F). The expression level of each RNA transcript was presented as 2-ΔCT (ΔCT is the difference of CT value between the specific RNA and GAPDH). The fold change of the specific RNA transcript after treatment was calculated using the formula: Fold change=2-ΔCT (ΔΔCT is the difference between the ΔCT of treated samples and that of control samples).
26 Measurement of ATP1A1-AS1 subcellular distribution
Nuclear and cytoplasmic RNA were extracted from cultured HK2 cells using a
RNA Subcellular Isolation Kit from Active Motif (Cat. No.: 25501). Extracted RNA was then subjected to reverse transcription for cDNA synthesis and RT-qPCR measurement using the same primer pairs as described above. Relative expression level in nuclear or cytosol was calibrated according to the total RNA amount obtained from each fraction.
Overexpression of ATP1A1-AS1 and FOXA1 in HK2 cells
Full length cDNA sequence of ATP1A1-AS1-203, Intron 2, or FOXA1 was cloned into a pcDNA3.1(-B) plasmid vector (Kumarasamy, Waghulde et al. 2015) and was verified by DNA sequencing. About 2 g per well of the overexpression vector was transfected into HK2 cells cultured in a 6-well plate with X-tremeGENE™ HP DNA
Transfection Reagent from Roche (Cat. No.: 6366236001) for 24 or 48 h. The same amount of empty pcDNA3.1(-B) vector was used as control.
Western blot
Control or treated HK2 cells were washed with ice cold phosphate-buffered saline
(PBS) once and solubilized in Radioimmunoprecipitation assay (RIPA) buffer containing
2mM PMSF, 1% protease inhibitor cocktail, and 1mM sodium orthovanadate from Santa
Cruz biotechnology (Cat. No.: sc-24948). After centrifuged at 14,000g for 15 minutes, the supernatants were collected and used for Western blot. The primary antibodies used in western blot analyses were: anti-Na/K ATPase α1 antibody (Developmental Studies
Hybridoma Bank at the University of Iowa, Cat. No.: α6F); anti-GAPDH antibody (Santa
Cruz Biotechnology, Cat. No.: sc-25778); anti-phospho-Src (pTyr418) antibody (Sigma-
27 Aldrich, Cat. No.: S1940); and anti-c-Src antibody (Santa Cruz Biotechnology, Cat. No.: sc-8056).
Cell proliferation assay
For cell proliferation assay, about 20,000 HK2 cells were seeded in each well on a
12-well plate and cultured for 24h. The cells were then transfected with pcDNA3.1(-B) vector containing ATP1A1-AS1-203. Cells transfected with empty pcDNA3.1(-B) vector were used as control. To monitor the cell proliferation, transfected cells with or without ouabain treatment were incubated for 96 h in an IncuCyte® S3 Live Cell Analysis System
(Essen BioScience) equipped with a microscope camera that automatically taking pictures from 9 different regions of each well every 2 h. Cell confluences were calculated and analyzed using the IncuCtye S3 live cell analysis software provided by the manufacturer.
Statistics
The data are presented as the Mean ± SEM and analyzed using Two-way ANOVA or Student T-test where appropriate. A p-value<0.05 is considered as significant.
28 Funding and Disclosure
Funding: This research was funded by NIH HL105649 to J.T. and was funded by
AHA 16SDG27700030 to S.K.
Author Contributions: Conceptualization, J.T. and S.K.; Methodology, J.T., X.
F., U.M.A, C.A.D., H.S., and S.K.; Formal Analysis, X. F., X. Z., and J.T.; Data Curation,
X. F. and J.T.; Writing-Original Draft Preparation, X.F. and J.T.; Writing-Review
&Editing, J.T. and S.K.; Funding Acquisition, J.T. and S.K.
Conflicts of Interest: The authors declare no conflict of interest.
Abbreviations
lncRNAs Long non-coding RNAs
SAHA Suberoylanilide hydroxamic acid
HDAC Histone deacetylase
CTS Cardiotonic steroids
29 Figure legends
Figure 1. Schematic presentation of ATP1A1 and ATP1A1-AS1 gene. Sequence information was obtained from Ensembl (GRCh38).
Figure 2. Differential expression and subcellular distribution of ATP1A1-AS1 splice variants in human kidney cells. (A): Expression of four splice variants of ATP1A1-AS1 and mRNA level of ATP1A1 in HK2 cells. (B): Expression of ATP1A1-AS1 splice variants and ATP1A1 in HEK293 cells. (C): Expression of ATP1A1-AS1 splice variants in cytosol and nuclear fraction of HK2 cells. A commercial isolation kit was used to extract nuclear and cytosol fractions. These experiments were repeated 4 times.
Figure 3. Epigenetic regulation of ATP1A1-AS1 expression in HK2 cells. Cultured
HK2 cells were treated with Deacetylase inhibitor SAHA (A) or DNA methylation inhibitor Decitabine (B) for 48 h and the total RNA was extracted using a commercial RNA extraction kit as described in Method section. Solvent-treated cells were used as control.
Expression of ATP1A1-AS1-203 was quantified using RT-qPCR. Data was presented as
Mean ± SEM and analyzed using Two-way ANOVA.
Figure 4. The effect of FOXA1 on ATP1A1-AS1 expression in HK2 cells. A pcDNA3.1(-B) plasmid that overexpresses FOXA1 (FOXA1) or empty vector (EV) was transfected into the cultured HK2 cells for 48 h, an empty pcDNA3.1(-B) plasmid was used as control. The expression of ATP1A1-AS1-203 (A) and ATP1A1 (B) was measured from the cell lysates using RT-qPCR. The fold change of the gene expression was calculated using CT method. The experiments were repeated 3 times.
30 Figure 5. Overexpression of ATP1A1-AS1-203 negatively regulates ATP1A1 gene expression and Na/K-ATPase α1 protein level in HK2 cells. HK2 cells were transfected with pcDNA3.1(-B) plasmid that overexpresses ATP1A1-AS1-203 (AS3) or empty vector
(EV) for 48 h and expression of ATP1A1-AS1-203 (A) and ATP1A1 (B) was measured using the RNA extracted from the cell lysate. The Na/K-ATPase α1 protein level (C) was measured by Western blot using the RIPA buffer resolved cell lysates.
Figure 6. Effect of intron 2 of ATP1A1-AS1-203 on the expression of ATP1A1 in HK2 cells. (A): The expression level of ATP1A1-AS1-203 and ATP1A1-AS1-203-Intron 2 in
HK2 cells. (B & C): The effect of ATP1A1-AS1-203-Intron 2 overexpression on ATP1A1 gene expression. The ATP1A1-AS1-203-Intron 2 overexpression vector (Intron2) was transient transfected into cultured HK2 cells for 48 h and the total RNA extracted from the cell lysates were used for RT-qPCR measurement of ATP1A1-AS1-203-Intron 2 (B) and
ATP1A1 (C). Experiments were repeated 4 times.
Figure 7. Overexpression of ATP1A1-AS1-203 inhibits ouabain-induced Src activation in HK2 cells. HK2 cells were transfected with ATP1A1-AS1-203 overexpressing vector (AS3) or empty vector (EV) for 24 h followed by ouabain (Oua) treatment for 15min. Cell lysates were collected in RIPA buffer. The phospho-Src at
Tyr418 (pSrc418) and c-Src were probed using Western blot. The upper panel was a representative Western blot, and the lower panel was the quantification result. The experiment was repeated 4 times and Two-way ANOVA was used for statistical analysis.
Figure 8. Overexpression of ATP1A1-AS1-203 inhibits HK2 cell growth in presence or absence of ouabain. HK2 cells were transfected with ATP1A1-AS1-203 vector (AS3)
31 or empty vector (EV) for 48 h followed by treatment of 50 nM ouabain (Oua) for another
48 h. Cells without ouabain treatment was used as control. Cell proliferation was monitored using an Incucyte incubator equipped with a camera that automatically taking pictures every 2 hours at 9 different regions of each cell culture well. (A): Cell growth curve in AS3 or EV transfected HK2 cells without ouabain treatment. (B): Cell growth curve in AS3 or
EV transfected HK2 cells that treated with 50 nM ouabain. Each experiment was repeated
4 times.
32 Figures
Figure 1. Schematic presentation of ATP1A1 and ATP1A1-AS1 gene.
33
Figure 2. Differential expression and subcellular distribution of ATP1A1-AS1 splice variants in human kidney cells.
34
Figure 3. Epigenetic regulation of ATP1A1-AS1 expression in HK2 cells.
35
Figure 4. The effect of FOXA1 on ATP1A1-AS1 expression in HK2 cells.
36
Figure 5. Overexpression of ATP1A1-AS1-203 negatively regulates ATP1A1 gene expression and Na/K-ATPase α1 protein level in HK2 cells.
37
Figure 6. Effect of intron 2 of ATP1A1-AS1-203 on the expression of ATP1A1 in HK2 cells.
38
Figure 7. Overexpression of ATP1A1-AS1-203 inhibits ouabain-induced Src activation in HK2 cells.
39
Figure 8. Overexpression of ATP1A1-AS1-203 inhibits HK2 cell growth in presence or absence of ouabain.
40 Chapter 4
Na/K-ATPase Signaling Mediates miR-29b-3p Regulation and Cardiac Fibrosis Formation in Mice with Chronic Kidney Disease
Christopher A. Drummond1*, Xiaoming Fan1, Steven T. Haller1, David J.
Kennedy1, Jiang Liu2, and Jiang Tian1, #
1 Department of Medicine at the University of Toledo, Toledo, OH 43614, United
States of America
* Current address: MPI Research, Mattawan, MI 49071, United States of America
2 Joan C. Edwards School of Medicine, Marshall University, Huntington, WV
25701, United States of America
Short Title: Na/K-ATPase on miR-29b-3p and Cardiac Fibrosis
# Address of Corresponding Author:
Jiang Tian, Ph.D.
Associate Professor
Department of Medicine
University of Toledo 41 3000 Arlington Ave.
Toledo, OH
Tel.: (419) 383-3510;
E-mail: [email protected]
This work has been published in PLoS One 13(5): e0197688.
42 Abstract
The Na/K-ATPase is an important membrane ion transporter and a signaling receptor that is essential for maintaining normal cell function. The current study examined the role of Na/K-ATPase signaling in regulating miR-29b-3p, an anti-fibrotic microRNA, in a mouse chronic kidney disease (CKD) model (5/6th partial nephrectomy or PNx). The results showed that CKD induced significant reduction of miR-29b-3p expression in the heart tissue by activation of Src and NFB signaling in these animals. To demonstrate the role of Na/K-ATPase signaling, we also performed the PNx surgery on Na/K-ATPase α1 heterozygous (α1+/-) mice, which expresses ~40% less Na/K-ATPase α1 compared to their wild type littermates (WT) and exhibits deficiency in Na/K-ATPase signaling. We found that CKD did not significantly change the miR-29b-3p expression in heart tissue from the
α1+/- animals. We also found that CKD failed to activate Src and NFB signaling in these animals. Using isolated cardiac fibroblasts from α1+/- mice and their WT littermates, we showed that ouabain, a specific Na/K-ATPase ligand, induces decreased miR-29b-3p expression in fibroblasts isolated from WT mice, but had no effect in cells from α1+/- mice.
Inhibition of NFB by Bay11-7082 prevented ouabain-induced miR-29b-3p reduction in
WT fibroblasts. To further confirm the in vivo effect of Na/K-ATPase signaling in regulation of miR-29b-3p and cardiac fibrosis in CKD animals, we used pNaKtide, a Src inhibiting peptide derived from the sequence of Na/K-ATPase, to block the activation of
Na/K-ATPase signaling. The result showed that pNaKtide injection significantly increased miR-29b-3p expression and mitigated the CKD-induced cardiac fibrosis in these animals.
These results clearly demonstrated that Na/K-ATPase signaling is an important mediator in CKD that regulates miR-29b-3p expression and cardiac fibrosis, which provides a novel
43 target for regulation of miR-29b-3p in CKD. We also demonstrate that antagonizing Na/K-
ATPase signaling by pNaKtide can reduce organ fibrosis through the stimulation of tissue miR-29b-3p expression.
Key Words: Na/K-ATPase; microRNA 29b; pNaKtide; Fibrosis; Chronic Kidney
Disease.
44 Introduction
MicroRNAs (miRNAs) are a group of non-coding RNAs that are 18-25 bp in length. These small RNAs can bind to the targeted mRNAs and inhibit their translation into functional proteins (Filipowicz, Bhattacharyya et al. 2008, Bartel 2009, Jiang, Tsitsiou et al. 2010). Studies have found a series of miRNAs that specifically target the mRNAs of extracellular matrix proteins and are associated with tissue fibrosis (Ambros 2004, van
Rooij, Sutherland et al. 2008, Jiang, Tsitsiou et al. 2010, Latronico and Condorelli 2011,
Qin, Chung et al. 2011, Noetel, Kwiecinski et al. 2012, Zhang, Huang et al. 2014).
Regulation of miRNAs is important in therapeutic interventions, and strategies have been developed and tested in several disease settings including cardiovascular disease (Li, Yong et al. 2010, Montgomery and van Rooij 2010, Bai, Xu et al. 2011, Shin, Jin et al. 2011,
Rathore, Saumet et al. 2012, Zhang, Fei et al. 2012). Studies have shown that reduction of miR-29b-3p can cause fibrosis in heart, lung, liver, skin and kidney, while increase of miR-
29b-3p prevents tissue fibrosis (van Rooij, Sutherland et al. 2008, Qin, Chung et al. 2011,
Wang, Kwan et al. 2012, Ramdas, McBride et al. 2013, Villa, Ridolfi et al. 2013, Zhu,
Chen et al. 2013). Cardiac fibrosis is common in cardiac diseases, and accumulation of collagen and other matrix proteins in the extracellular space forms interstitial fibrosis (Last
1985, Brecher, Hinko et al. 1996, Goldsmith, Bradshaw et al. 2013, Kong, Christia et al.
2014). Experimental and clinical data showed that fibrosis increases cardiac stiffness, but reduction of fibrosis may improve cardiac function (Jalil, Doering et al. 1989, Brilla, Funck et al. 2000). Fibrosis could also lead to sudden cardiac death even in conditions that has no cardiac symptoms (Gulati, Jabbour et al. 2013).
45 Na/K-ATPase is an important cell membrane protein enriched in heart and kidney tissues. The Na/K-ATPase signaling involves Src, Akt, PKC, and other downstream signaling proteins (Xie and Askari 2002). Our laboratory has previously shown that formation of cardiac fibrosis involves increases in endogenous cardiotonic steroids (CTS) and activation of Na/K-ATPase signaling in the 5/6th partial nephrectomy (PNx) model of chronic kidney disease (CKD) (Kennedy, Vetteth et al. 2006, Kennedy, Elkareh et al.
2008, Haller, Kennedy et al. 2012). We also found that in isolated cardiac fibroblasts, treatment with ouabain, a Na/K-ATPase ligand, induced decreases in miR-29b-3p and increased collagen expression, whereas blocking the Na/K-ATPase related signaling pathway restores miR-29b-3p levels and mitigated collagen expression in these cells
(Drummond, Hill et al. 2016). The current study was designed to test the in vivo effect of
Na/K-ATPase signaling in regulating the expression of miR-29b-3p and tissue fibrosis in a mouse model of CKD.
Results
Na/K-ATPase is Involved in the Regulation of miR-29b-3p and Cardiac
Fibrosis in CKD Animals.
To test the role of the Na/K-ATPase in CKD animals, we performed PNx surgery on WT mice as well as on α1+/- mice. Cardiac tissue was collected at 16 weeks after PNx surgery. As shown in Figure 1A, cardiac Na/K-ATPase α1 subunit expression in 1+/- mice was ~40% less than that in WT mice, which is consistent with previous reports
(Moseley, Cougnon et al. 2004, Liu, Bai et al. 2012). When miR-29b-3p was measured by
RT-qPCR using total RNA extracted from left ventricle tissue, we found that PNx
46 decreased miR-29b-3p levels by about 2 fold in WT mice (0.89±0.14 in sham vs 0.46±0.08 in PNx, p<0.01), whereas in α1+/- mice the basal level of miR-29b-3p was slightly lower than that in WT mice, but PNx surgery caused no significant changes in miR-29b-3p expression (Figure 1B). We also measured the potential targets of miR-29b-3p such as collagen 1A1, matrix metalloproteinase-2 (Mmp-2), fibrillin 1 (Fbn1), and elastin (Eln).
As shown in Fig 1C, PNx significantly increased collagen 1A1 and Mmp-2 by ~50% and
100%, respectively, in WT mice, but had no significant effect in α1+/- mice. The expression of Fbn1 and Eln was not significantly changed in either WT or α1+/- mice.
Na/K-ATPase Regulates miR-29b-3p through Activation of Src and NFB in
CKD mice.
It has been previously reported that Na/K-ATPase regulates Src (Kennedy, Elkareh et al. 2008, Liu, Bai et al. 2012) and NFB signaling (Li, Zelenin et al. 2006, de Sa Lima,
Kawamoto et al. 2013), while both Src and NFB are known to regulate miR-29b-3p expression (Liu, Wu et al. 2010, Drummond, Hill et al. 2016). We then examined these signaling pathways in both WT and α1+/- mice. As shown in Figure 2A, PNx surgery significantly increased Src phosphorylation at Tyr418 (an indication of Src activation) in the left ventricle tissue from WT mice, while it failed to induce activation of Src in α1+/- mice, albeit the basal level in α1+/- mice is higher compared to that in WT mice. PNx also caused higher expression of total NFB p65 (Figure 2B) as well as its nuclear fraction
(Figure 2C) in the left ventricle tissue from WT mice. Similarly, the basal level of NFB p65 is higher in α1+/- mice, but PNx did not further increase the expression level or nuclear fraction of NFB.
47 We also performed immunostaining for NFB using an anti-NFB p65 antibody.
As shown in Figure 3A, the NFB p65 signal was weak in sham-operated WT mice, but
PNx caused a significant increase in NFB levels, especially in the smaller cells between myocytes. Most of the NFB signal (green) was around or colocalized with the DAPI signal (purple) in these cells, indicating a nuclear transfer of NFB p65. However, in sham- operated 1+/- mice (Figure 3B), the NFB p65 signal in the left ventricle tissue was much higher than that in WT mice. Interestingly, in the α1+/-mice, the NFB staining was in a punctate pattern and mostly in myocytes. PNx surgery caused no significant change in
NFB p65 expression or its location in α1+/- mice. These results of NFB expression and activation are consistent with the observation of miR-29b-3p changes in WT and α1+/- mice, suggesting that activation of Src and NFB may mediate the PNx-induced reduction of miR-29b-3p in WT mice.
We have previously demonstrated that Src activation is involved in Na/K-ATPase regulated miR-29b-3p in isolated cardiac fibroblasts (Drummond, Hill et al. 2016). To further determine the role of NFB in Na/K-ATPase mediated miR-29b-3p regulation, we used isolated cardiac fibroblasts from WT as well as α1+/- mice. These fibroblasts were treated with ouabain (a specific Na/K-ATPase ligand) alone or in combination with a specific NFB inhibitor (Bay11-7082) for 24h. Total RNA was extracted from the cell lysates and miR-29b-3p level was quantified using RT-qPCR as described previously
(Drummond, Hill et al. 2016). As shown in Figure 4A, ouabain alone induced a decrease in miR-29b-3p expression in a dose-dependent manner in fibroblasts isolated from WT mice and the decrease of miR-29b-3p can be blocked by Bay11-7082, suggesting that
NFB is involved in Na/K-ATPase mediated miR-29b-3p regulation. However, in cardiac
48 fibroblasts isolated from α1+/- mice, ouabain failed to induce the change in miR-29b-3p levels, and the NFB inhibitor had no effect either (Figure 4B).
Treatment with pNaKtide inhibits Src Activation and Antagonizes PNx-
Induced Cardiac Fibrosis by Increasing miR-29b-3p Expression in Heart Tissue from
CKD Mice.
The pNaKtide has been reported as an inhibitor of Na/K-ATPase related Src signaling (Li, Cai et al. 2009, Li, Zhang et al. 2011), which prevents ouabain-induced reduction of miR-29b-3p in isolated cardiac fibroblasts (Drummond, Hill et al. 2016). To test the effect of pNaKtide on miR-29b-3p expression and cardiac fibrosis in CKD mice, we performed PNx or sham surgery on WT mice and α1+/- mice. At 12 weeks after the
PNx or sham surgery, pNaKtide was given by intraperitoneal injection at 25 mg/kg bodyweight every other week for a total of 3 injections. Mice injected with the same volume of saline were used as control. Organ collection was done at the end of the 16th week after surgery. As shown in Figure 5, PNx alone caused higher Src phosphorylation at
Tyr418 in left ventricle tissue from WT mice, while pNaKtide injection attenuated the
PNx-induced Src phosphorylation. In α1+/- mice, the basal level of Src phosphorylation was higher compared to that in WT, but PNx was unable to further stimulate Src phosphorylation and actually decreased Src phosphorylation in these animals. Injection of pNaKtide did not change the phosphorylation state compared to PNx alone in α1+/- mice.
Consistently, as shown in Figure 6, we found that injection of pNaKtide in WT mice significantly increased miR-29b-3p expression in heart tissue compared to that in PNx alone group. In α1+/- mice, pNaKtide slightly increased miR-29b-3p expression in heart
49 tissue. Injection of pNaKtide in sham-operated mice caused a slight increase in miR-29b-
3p expression, but it was not statistically significant.
To test if the above changes in Src activation and miR-29b-3p expression correlate with cardiac tissue fibrosis, formalin-fixed left ventricle tissue was analyzed using
Trichrome staining as described in the Materials and Methods section. As shown in Figure
7, PNx significantly increased cardiac fibrosis by about 9-fold in WT mice (0.60% in sham vs 5.3% in PNx, p<0.01), while treatment with pNaKtide led to significantly reduced fibrosis compared to PNx alone (1.3 ± 0.3% in the PNx plus pNaKtide group vs 5.3 ± 1.5 in the PNx alone group, p<0.01) in WT mice. In α1+/- mice, the basal level of cardiac fibrosis was higher than that in WT mice, and PNx induced a milder 3-fold change in cardiac fibrosis (0.8±0.2% in sham vs 2.3±0.6% in PNx, p<0.05). Injection of pNaKtide also reduced cardiac fibrosis in α1+/- mice (0.7±0.2% in PNx plus pNaKtide group vs
2.3±0.6% in PNx alone), but did not reach statistical significance. Treatment with pNaKtide in sham-operated mice yielded no significant changes in levels of cardiac fibrosis.
Treatment with pNaKtide Ameliorates PNx-induced Cardiac Hypertrophy.
In addition to its effects on fibrosis, we also examined the effect of pNaKtide on
PNx-induced cardiac hypertrophy. As shown in Figure 8A, the heart weight/body weight
(HW/BW) ratio of animals subjected to PNx was significantly increased compared to sham-operated animals at 16 weeks (6.20.7 in PNx group vs 4.20.2 in sham group, p<0.01). Injection of pNaKtide led to a 23.3% reduction of cardiac mass compared to PNx alone animals. In α1+/- mice, PNx also increased cardiac hypertrophy, but to a much
50 smaller extent (4.80.2 in PNx vs 4.20.2 in sham, p>0.05). Consistent with these observations, as shown in Figure 8B, we found that PNx led to more enlarged cardiac myocytes in WT animals when the sizes of individual cardiac myocytes were analyzed using Wheat Germ Agglutinin (WGA) staining. The average cross sectional area was significantly reduced with pNaKtide injection in WT mice (461 ± 13 nm2 in PNx alone vs
386 ± 10 nm2 in pNaKtide injected animals, p<0.05). These effects of PNx or pNaKtide injection in α1+/- mice were much less pronounced.
We also evaluated other cardiac function using echocardiography. The echo data were summarized in Tables 1 and 2. We found that in WT animals there was a significant increase in diastolic dimension at 16 weeks after PNx surgery, and pNaKtide injection attenuated the increase in diastolic dimension. Ejection fraction (EF) was significantly decreased with PNx in both WT and α1+/- mice and injection of pNaKtide slightly improved EF in WT but not in α1 +/- mice.
Discussion
Multiple studies have found a significant role of anti-fibrotic microRNAs in the prevention of cardiac fibrosis (Ambros 2004, van Rooij, Sutherland et al. 2008, Jiang,
Tsitsiou et al. 2010, Montgomery and van Rooij 2010, Latronico and Condorelli 2011,
Noetel, Kwiecinski et al. 2012). Our previous data showed that prolonged activation of
Na/K-ATPase signaling contributes to cardiac fibrosis in uremic cardiomyopathy (Elkareh,
Periyasamy et al. 2009). More recently, we found that mimicry of miR-29b-3p can prevent
CTS-induced collagen synthesis in cardiac fibroblasts (Drummond, Hill et al. 2016),
51 indicating that an increase in miR-29b-3p expression is a potential therapeutic treatment strategy for fibrosis-related diseases. In addition, our in vitro data showed that pNaKtide treatment prevented excessive collagen synthesis by restoring endogenous miR-29b-3p expression to levels close to non-treated controls (Drummond, Hill et al. 2016). The current study further showed the same effect of pNaKtide in vivo as well as its potential in attenuating PNx-induced cardiac fibrosis and hypertrophy.
Mechanistically, the results from this study demonstrate that CKD induces activation of Na/K-ATPase-mediated Src and its downstream target NFB. As a transcription factor, NFB can form a complex with Sp1 and HDAC and directly bind to the regulatory sequence of the miR-29b-3p gene, causing decreased expression of miR-
29b-3p (Liu, Wu et al. 2010). Disrupting the Na/K-ATPase-related signaling and inhibition of Src activation by pNaKtide increased miR-29b-3p expression in heart tissue and thus attenuated cardiac fibrosis in these animals. The data from α1+/- mice also demonstrated that miR-29b-3p regulation requires Na/K-ATPase signaling activation. Reduction of
Na/K-ATPase α1 caused a deficiency in Src and NFB activation, and CKD caused no significant changes on miR-29b-3p expression in these animals. However, an interesting observation is that even though the basal level of Src phosphorylation and NFB activation is higher in heart tissue from α1+/- mice compared to their WT littermates, the miR-29b-
3p expression was not significantly different between these animals. The specific mechanism for this phenomenon is still elusive. Our previous studies also showed that activation of Src by ouabain treatment in pig kidney proximal tubule cells induced cell proliferation, whereas in Na/K-ATPase-reduced cell lines ouabain caused cell growth inhibition or death despite the increased basal level of Src phosphorylation in these cells
52 (Tian, Li et al. 2009). Similarly, we demonstrated that the increased basal level of Src phosphorylation cannot prevent cardiac cell apoptosis in α1+/- mice when infused with marinobufagenin, another specific Na/K-ATPase ligand (Liu, Bai et al. 2012). A possible explanation is that Na/K-ATPase reduction-induced Src activation is an adaptive process.
This adaptive change in Src phosphorylation is required to maintain the normal cellular function, but it cannot further respond to extrinsic stress such as CKD or ouabain treatment.
It could also be due to the compartmentalization of these signaling molecules.
Reduction of Na/K-ATPase in α1+/- mice may also cause the change in Na+ and
K+ transporting activity, especially in the kidney, which only expresses the α1 subunit of
Na/K-ATPase. However, the measurement of plasma Na+ concentration (148.8±0.9 mM, n=5) from these animals was not significantly different from WT animals (148.4±1.9 mM, n=5), suggesting that the ion transporting activity may be compensated by increased per unit activity of Na/K-ATPase. This is in line with our previous observation that reduction of Na/K-ATPase α1 subunit by 40% resulted in only about 20% decrease in ion transporting activity in pig kidney epithelial cell line (Liang, Tian et al. 2007).
In addition, we found that even though Na/K-ATPase reduction in α1+/- mice caused a deficiency in Na/K-ATPase signaling and prevented miR-29b-3p dysregulation, it did not completely block PNx-induced cardiac fibrosis, suggesting that other pathways also contribute to the formation of cardiac fibrosis in this model. Our previous studies showed that cardiac apoptosis increased in α1+/- mice subjected to PNx surgery or MBG infusion (Liu, Bai et al. 2012, Drummond, Sayed et al. 2014). These apoptotic events may cause the release of cytokines that stimulate fibrotic changes in the tissue. These results also suggest that the reduction of Na/K-ATPase may not be an appropriate strategy to
53 reduce CKD-related cardiac fibrosis. In fact, a decrease in Na/K-ATPase has been reported in heart failure patients and the amount of Na/K-ATPase was correlated with the ejection fraction in these patients (Norgaard, Bagger et al. 1988, Ishino, Botker et al. 1999).
In summary, the current study demonstrates that Na/K-ATPase signaling is an important mediator that regulates miR-29b-3p, which contributes to the formation of cardiac fibrosis in the setting of CKD.
Materials and Methods
Animals: Animal experiments were conducted in accordance with the National
Institutes of Health, Guide for the Care and Use of Laboratory Animals under the protocol
(IACUC# 106846) approved by the Institutional Animal Care and Use Committee at the
University of Toledo. Na/K-ATPase α1 subunit heterozygous (α1+/-) and wild type (WT) mice were generated from C57/Black Swiss mice as previously described (James, Grupp et al. 1999). These mice were obtained from Dr. Jerry Lingrel at the University of
Cincinnati and maintained in our animal facility. The heterozygous and WT mice were crossed to generate the inbred WT and heterozygous offspring that were used for these experiments. The heart tissue of α1+/- mice contains ~40% less Na/K-ATPase 1 compared to their WT littermates (Moseley, Cougnon et al. 2004, Liu, Bai et al. 2012).
Adult male mice at 2-3 months of age and weighing between 25-27g were used for this study. All mice were reared under a 12h dark/light cycle, fed standard chow and were provided water ad libitum. These conditions were utilized for the entire duration of the experiment.
54 Male α1+/- mice and their WT littermates were each divided into two groups based on surgical intervention: the first group consisted of sham-operated animals as controls; the second group of animals were subjected to 5/6th partial nephrectomy (PNx) surgery.
Following the surgery, each group was further divided into two subgroups: one group received intraperitoneal injection of pNaKtide at the 12th week post-surgery at a dose of
25mg/kg bodyweight, and another group received the same volume of saline as control.
PNx surgery was performed as previously described (Drummond, Sayed et al. 2014).
Briefly, mice were anesthetized with a mixture of 100% oxygen and 2% isoflurane. An incision was made in the left flank on the back, the left kidney was exposed, and the artery supplying the upper pole of the kidney was ligated with a 6-0 silk suture (Coviden,
Mansfield, Ma, Cat No: S-1750K) under a high-power dissecting microscope. The kidney was then reinserted to the body cavity and the incision was closed. One week later, the right kidney was surgically removed but the renal capsule with the adrenal gland were kept.
To alleviate the animals from possible pain due to the surgery, we subcutaneously injected a long-lasting buprenorphine (0.05mg/kg) at 30 min before the surgery.
Echocardiographic imaging, Blood Pressure Measurement, and Organ
Collection: Echocardiography was performed before surgery as baseline and at the end of the 16th week prior to organ collection using an Acuson Sequoia C512 machine (Siemens) as previously described (Drummond, Sayed et al. 2014). Briefly, animals were anesthetized with a mixture of 2% isoflurane and 100% oxygen and were secured to a heated metal platform in a supine position with medical tape on all four extremities. A 15 mHz linear transducer 15L8 (Siemens) was used to acquire images in a shallow left-side position.
55 Organ collection was done at the end of the 16th week following PNx surgery. Mice were euthanized by anesthesia with a mixture of ketamine and xylazine (100mg/kg and
10mg/kg, respectively) followed by exsanguination. One half of the heart left ventricle or the kidney was immediately placed in a 4% formaldehyde solution for fixation, while the other half was flash frozen in liquid nitrogen and stored at -80⁰C for later use for biochemical analysis.
Western Blot Analysis: Tissue homogenates were prepared by placing left ventricle tissue in ice-cold RIPA lysis buffer (pH 7.0) from Santa Cruz Biotechnology (Cat.
No.: SC-24948), followed by homogenization. Aliquots were then made for Western blot analysis or stored frozen at -80⁰C. Proteins from homogenates were separated by SDS- polyacrylamide gel electrophoresis. The resolved proteins were then electro-transferred to a Nitrocellulose membrane (Fisher Scientific, Hanover Park, IL; Cat. No. 45-004-007) for immunoblotting. Src kinase activation was determined using a primary antibody against phospho-Src at Tyr418 (Fisher Scientific, Cat. No.: 44660G; pSrc). Total Src was probed with a mouse anti-cSrc antibody from Santa Cruz Biotechnology (Cat. No.: SC-8056).
Expression of Na/K-ATPase α1 subunit was measured using an anti-Na/K-ATPase 1 antibody from the Developmental Studies Hybridoma Bank at the University of Iowa (Cat.
No.: α6F). GAPDH (Santa Cruz; Cat. No. SC-25778) was used as a loading control.
Histology: Left ventricle sections fixed in 4% formaldehyde solution (pH 7.2) were paraffin embedded and cut into a thickness of 4 μm onto microscopy slides. The tissue sections were deparaffinized with xylene and rehydrated by sequential incubations in
56 ethanol and water. Masson’s Trichrome staining for cardiac fibrosis was conducted on the
4 µm heart tissue sections. Computer aided morphometry was used to quantify the percent area of fibrosis as previously described (Kennedy, Elkareh et al. 2008, Drummond, Sayed et al. 2014). For each section, 10 images were randomly taken with a bright-field microscope with a 20X lens. The percentage of blue-color area was measured using ImageJ software. The average of the 10 images from each section was counted as one measurement for statistical analysis.
Wheat Germ Agglutinin (WGA) Staining: Paraffin-embedded left ventricle tissue sections (4 µm in thickness) were deparaffinized and rehydrated in four changes of
Xylene, 2 changes of 100% ethanol, 2 changes of 95% ethanol, and 2 changes of 70% ethanol. The tissue sections were then rinsed with tap water and incubated with Oregon
Green 488 WGA solution (5 µg/ml) from Life Technologies (Cat. No. W6748) at 4C overnight. Afterwards, slides were washed three times with PBS. The slides were allowed to air dry, and then mounted with Prolong Gold Antifade Reagent from Life Technologies
(Cat. No.: P36930). Eight fluorescent images were randomly taken from each tissue section using an Olympus fluorescent microscope with a 20x lens. From each image, the cross- sectional area of 20 cells was measured using ImageJ as previously described (Drummond,
Sayed et al. 2014).
Isolation, Culture and Treatment of Cardiac Fibroblasts: Isolation of cardiac fibroblasts was carried out as previously described (Elkareh, Kennedy et al. 2007,
Drummond, Hill et al. 2016). Briefly, hearts of adult male α1 +/- mice or their WT
57 littermates were anesthetized with Ketamine/Xylazine. The heart was cut off from the upper end of the ascending aorta and perfused under sterile conditions with Joklik’s medium from Sigma (Cat. No.: M0518) on a modified Langendorff Apparatus for 5 min.
Perfusate was switched to the Joklik’s medium containing 0.1% Collagenase (Sigma; Cat.
No.: C0130-1G) and 0.1% bovine serum albumin (BSA) for 25min. After perfusion the heart was cut into small pieces and shaken in the same medium for 30min at 37C with constant agitation. The resulting cell suspension was centrifuged at 600g for 10min, and the supernatant was centrifuged at 1500g for 15min. The resulting fibroblasts were cultured in Dulbecco’s Modified Eagle Medium from Sigma (Cat. No.: D1152) supplemented with antibiotics and 15% fetal bovine serum (Life Technologies Inc., Cat. No.: 10437-028). .
The second passage of cells was used for experiments.
Measurement of NFκB Expression and Activation: For total expression of
NFB measurement, left ventricle tissue homogenate was analyzed by Western blot using anti-NFB p65 antibody from Abcam (Cat. No. AB32536). To detect the nuclear fraction of NFB p65, left ventricle tissue was homogenized and the nuclear and cytoplasmic isolation was performed using a commercial extraction kit (Fisher Thermo, Cat. No.
78835). Western blot was then used to analyze the levels of NFB p65 in nuclear and cytosol fractions. Lamin B1 was used as a marker and loading control for the nuclear fraction. The Lamin B1 antibody was purchased from Abcam (Cat No: AB16048). The nuclear/cytosol ratio of NFB was calculated after correction by Lamin B1 and GAPDH, respectively.
58 Immunostaining for NFB P65 in Heart Tissue: Paraffin-embedded left ventricle tissue sections (4 µm in thickness) were deparaffinized as described above for
WGA staining. After blocking with 1% BSA for 1h at room temperature, slides were incubated with anti-NFB P65 antibody (Abcam, Cat. No: ab16502) at 4C overnight. The primary antibody was then washed with TBS-T solution for 3 times, followed by incubating with a secondary anti-rabbit antibody conjugated with Alexa 488 for 2h at room temperature. The slides were then incubated with mounting medium containing DAPI for nuclear staining and mounted with coverslip. Fluorescent signals were visualized using a
Leica confocal microscope with a 63x oil lens. Five images were taken from each slide.
Data from 4-5 animals in each group were analyzed by Two-Way ANOVA with GraphPad software version 7.0.
RNA Isolation and Reverse Transcription-Quantitative Polymerase Chain
Reaction (RT-qPCR): The left ventricular tissue was homogenized in Qiazol from Qiagen and total RNA was isolated using the miRNeasy mini kit following the protocol provided by the manufacturer. Approximately 250 ng of extracted total RNA was used to synthesize cDNA in the miScript II RT kit (Qiagen, Inc.; Cat. No.: 218160). Quantification of miR-
29b-3p expression was performed as previously described (Jones, Stroud et al. 2011,
Drummond, Hill et al. 2016). Briefly, Qiagen miScript primer for miR-29b-3p (Cat. No.:
MS00005936) was mixed with the cDNA and miScript SYBR Green solution provided in a commercial PCR kit (Cat. No.: 218161, Qiagen, Inc.). RT-PCR was performed on a
Qiagen Rotor-Gene Q PCR machine. Calculation of miRNA expression was conducted by comparing the relative change in cycle threshold value (ΔCt) between miR-29-3p and the
59 internal control, RNU6 (Cat No.: MS00033740 from Qiagen). Fold change in expression was calculated for each miRNA using the equation of 2-ΔΔCt.
Statistical Analyses: Data are presented as Mean ±Standard Error of the Mean
(SEM) and analyzed using One-way or Two-way ANOVA where they are appropriate.
60 Funding and Disclosure
* This work was supported by the National Institutes of Health (HL-105649 to JT,
HL-137004 to DJK, DK-106666 to JL, and F32DK104615 to CAD). Additional support was from the National Affiliate of the American Heart Association (12SDG12050473 to
DJK), the David and Helen Boone Foundation Research Fund (DJK), an Early Career
Development Award from the Central Society for Clinical and Translational Research
(DJK), the University of Toledo Women and Philanthropy Genetic Analysis
Instrumentation Center (DJK and STH), and the University of Toledo Medical Research
Society (STH).
We also thank Dr. Jerry B. Lingrel for providing us the wild type and Na/K-ATPase alpha 1 subunit heterozygous knockout mice.
61 Acknowledgement
Contributions:
Author Contributions: conceptualization: JT
Author Contributions: methodology: CAD, JT
Author Contributions: software:
Author Contributions: validation: JT, CAD
Author Contributions: formal analysis: CAD, XF, JT
Author Contributions: investigation: CAD, XF, JT
Author Contributions: resources: CAD, XF, JT
Author Contributions: data curation:
Author Contributions: writing (original draft preparation): JT, CAD
Author Contributions: writing (review and editing): STH, DJK
Author Contributions: visualization:
Author Contributions: supervision: JT
Author Contributions: project administration: JT
Author Contributions: Funding acquisition: JT
62 Tables
Table 1: Echocardiographic data in WT mice 16 weeks following PNx or Sham
Surgery
Sham ShamP PNx PNxP Variable (n=10) (n=6) (n=7) (n=9)
Heart Rate (BPM) 461 ± 14 511 ± 22 438 ± 12 458 ± 19
EDV (μL) 37.2 ± 2.2 33.6 ± 4.1 52.0 ± 4.5* 47.3 ± 4.3
ESV (μL) 18.1 ± 2.4 17.4 ± 2.8 33.0 ± 5.0* 24.7 ± 4.5
EF (%) 61.5 ± 5.0 48.3 ± 4.4 40.8 ± 4.6* 50.0 ± 3.0
IVSWT at Diastole (mm) 0.93 ± 0.07 0.86 ± 0.08 0.91 ± 0.09 0.92 ± 0.10
PWT at Diastole (mm) 0.94 ± 0.06 0.85 ± 0.08 0.78 ± 0.12 1.00 ± 0.10
LVMI 4.1 ± 0.3 3.4 ± 0.2 5.3 ± 0.9 4.6 ± 1.0
IVSWT: interventricular septal wall thickness; PWT: posterior wall thickness;
WTI: wall thickness index ([PWT + IVSWT]/DD). Left ventricle mass index (LVMI). Data presented as Mean ±SEM. *, indicates p<0.05 different from sham. Sham: sham-operated control group; ShamP: sham-operated group that received pNaKtide injection; PNx: mice subjected to PNx surgery; PNxP: mice subjected to PNx surgery and received pNaKtide injection.
63 Table 2: Echocardiographic data in 1+/- mice 16 weeks following PNx or
Sham Surgery
Sham ShamP PNx PNxP Variable (n=11) (n=5) (n=11) (n=7)
Heart Rate (BPM) 436±52 504±26* 458±34 472±28
EDV (μL) 31.6±2.7 40.1±5.6 43.8±2.4* 39.0±4.4
ESV (μL) 14.1±1.8 18.9±3.5 23.3±1.8* 23.2±3.3
EF (%) 56.5±2.8 53.7±3.5 47.1±2.6 41.9±3.6*
IVSWT at Diastole (mm) 0.86±0.05 0.75 ± 0.05 0.84 ± 0.05 0.93 ± 0.07
PWT at Diastole (cm) 0.85±0.04 0.76 ± 0.10 0.81 ± 0.04 0.88 ± 0.05
LVMI 3.2 ±0.2 3.2 ± 0.6 4.0 ± 0.3 3.9 ± 0.3
IVSWT: interventricular septal wall thickness; PWT: posterior wall thickness;
WTI: wall thickness index ([PWT + IVSWT]/DD). Left ventricle mass index (LVMI). Data presented as Mean ±SEM. *, indicates p<0.05 different from sham. Sham: sham-operated control group; ShamP: sham-operated group that received pNaKtide injection; PNx: mice subjected to PNx surgery; PNxP: mice subjected to PNx surgery and received pNaKtide injection.
64 Figure Legends
Figure 1. PNx-induced miR-29b-3p expression change in WT and α1+/- mice.
WT and 1+/- mice were subjected to sham or 5/6th partial nephrectomy (PNx) surgery and left ventricle tissue was collected at the end of 16th week for RNA extraction, RT- qPCR, and Western blot analyses. A): Western blot of Na/K-ATPase 1 subunit expression in left ventricle tissue from different group of mice. !! indicates p<0.01 1+/- vs WT. B):
Expression of miR-29b-3p measured by RT-qPCR in left ventricle tissue from experimental mice. Data was analyzed using Two-Way ANOVA with GraphPad software
7.0., N=5 in each group * indicates p<0.05 PNx vs Sham. C): mRNA expression of miR-
29b-3p targeted genes: collagen 1A1 (Col1a1), matrix metalloproteinase-2 (Mmp2), fibrillin 1 (Fbn1), and elastin (Eln). * indicates p<0.05 PNx vs Sham; ** indicates p<0.01
PNx vs Sham. !! indicates p<0.01 1+/- vs WT.
Figure 2. Activation of Src and NFB in mice left ventricle tissue after PNx surgery. A): Western blot shows Src activation indicated by phosphorylation at Tyrosine
418 (pSrc) in left ventricle tissue, normalized to total Src levels (cSrc). Data are normalized as percentage expression of WT Sham animals. B): Western blots for total NFκB p65 in heart left ventricle tissue homogenates from each group. C): Western blots of NFκB p65 in nuclear and cytosolic fractions of isolated from left ventricle tissue. Lamin B1 is used as a nuclear marker. Nuclear/cytosolic ratio of NFB was calculated as indicator of NFB activation. Data were presented as mean ± SEM from 5 mice in each group. Data were analyzed using Two-Way ANOVA followed by Pairwise comparisons using Tukey’s correction for multiple comparisons, N=4 in each group. * indicates p<0.05 PNx vs sham;
65 ** indicates p<0.01 PNx vs Sham; !! indicates p<0.01 1+/- sham vs WT sham or α1+/-
PNx vs WT PNx.
Figure 3. Expression and translocation of NFB in heart tissue after PNx surgery. Left ventricle tissue slides (4 µm in thickness) from WT sham (N=4), WT PNx
(N=5), α1+/- sham (N=4), and α1+/- PNx (N=4) animals were deparaffinzed and immunostained for NFB p65. (A): Representative images from WT mice; (B):
Representative images from α1+/- mice; (C): Quantification data. Colocalization of NFB and DAPI was analyzed using ImageJ. The ratio of NFB p65 positive nuclear and total nuclear was used for analysis by Two-Way ANOVA. ** indicates p<0.01, WT PNx vs WT sham. Scale bar is 25 µm.
Figure 4. The effect of NFB inhibitor on ouabain-induced miR-29b-3p regulation in cardiac fibroblasts isolated from WT and α1+/- mice. Isolated primary cultures of mouse cardiac fibroblasts were pretreated with 1µM of the NFκB inhibitor BAY
11-7082 for 15 min followed by ouabain (10 or 100nM) treatment for 24h. Non-treated or ouabain treatment alone without BAY 11-7082 was used as control. After treatment, cell lysates were collected. Expression of miR-29b-3p was measured using RT-qPCR in RNA isolated from cells from WT animals (A) and α1 +/- animals (B). Fibroblasts were obtained from 4 animals in each group. Data was analyzed using One-Way ANOVA with GraphPad software 7.0. * indicates p<0.05 versus non-treated controls. ! indicates p<0.05 Bay11-
7082 plus ouabain vs ouabain alone.
Figure 5. Injection of pNaKtide blocks Src activation in mice subjected to PNx surgery. Tissue homogenate obtained from left ventricle tissue were used to probe phosphor-Src (pSrc) and total Src (cSrc) using Western blot. Data were obtained from 5
66 animals in each group and normalized as percentage expression of WT Sham and was analyzed using Two-Way ANOVA with GraphPad software 7.0. Sham: sham-operated animals, ShamP: sham-operated animals with pNaKtide injection. PNx: PNx-operated animals, PNxP: PNx-operated animals with pNaKtide injection.* indicates p<0.05 vs
Sham; & indicates p<0.05, PNxP vs PNx; !! indicates p<0.01 α1+/- PNx vs WT PNx.
Figure 6. Injection of pNaKtide diminishes PNx-induced decrease in miR-29b-
3p expression. Total RNA obtained from left ventricle tissue were used for RT-qPCR analyses. Expression of miR-29b-3p was presented as fold regulation relevant to WT sham animals. Data were obtained from 5 animals in each group and was analyzed using Two-
Way ANOVA with GraphPad software 7.0. * indicates p<0.05 vs Sham; & indicates p<0.05
PNxP vs PNx.
Figure 7. Injection of pNaKtide mitigates PNx-induced cardiac fibrosis. IP injection of 25mg/kg pNaKtide was performed biweekly started at 12th week following
PNx surgery and heart left ventricle tissue was collected at the end of 16th week. Cardiac fibrosis was evaluated using Trichrome staining. Data are presented as Mean ± SEM, N=10 for Sham WT, 6 for ShamP WT, 7 for PNx WT, 9 for PNxP WT, 11 for Sham α1+/-, 5 for
ShamP α1+/-, 11 for PNx α1+/-, 7 for PNxP α1+/. The upper panel are representative 20x
Trichrome staining. Scale bar =25μm. The lower panel is the quantification data of fibrosis
(% area). Data was analyzed using Two-Way ANOVA with GraphPad software 7.0. ** indicates p<0.01 vs Sham; & indicates p<0.05 PNxP vs PNx; !! indicates p<0.01 α1+/- PNx vs WT PNx.
Figure 8. Injection of pNaKtide attenuates PNx-induced cardiac hypertrophy.
A): Data of heart weight/body weight (HW/BW) ratio were obtained from 6-10 mice from
67 each group. B): Myocytes cross sectional area were evaluated using wheat germ agglutinin
(WGA) staining as described in Method section. Eight images were randomly taken from each tissue slide and 20 cells from each image were measured for their cross-sectional area.
Data is presented as Mean SEM and analyzed using Two-Way ANOVA with GraphPad software 7.0. * indicates p<0.05 vs Sham; ** indicates p<0.01 vs Sham; & indicates p<0.05
PNxP vs PNx.
68 Figures
Figure 1. PNx-induced miR-29b-3p expression change in WT and α1+/- mice.
69
Figure 2. Activation of Src and NFκB in mice left ventricle tissue after PNx surgery.
70
Figure 3. Expression and translocation of NFκB in heart tissue after PNx surgery.
71
Figure 4. The effect of NFκB inhibitor on ouabain-induced miR-29b-3p regulation in cardiac fibroblasts isolated from WT and α1+/- mice.
72
Figure 5. Injection of pNaKtide blocks Src activation in mice subjected to PNx surgery.
73
Figure 6. Injection of pNaKtide diminishes PNx-induced decrease in miR-29b-3p expression.
74
Figure 7. Injection of pNaKtide mitigates PNx-induced cardiac fibrosis.
75
Figure 8. Injection of pNaKtide attenuates PNx-induced cardiac hypertrophy.
76 Chapter 5
Summary
In this study, we characterized a natural antisense RNA, ATP1A1-AS1, that negatively regulates Na/K-ATPase α1 expression in human kidney cells. We further found that overexpression of ATP1A1-AS1 disproportionally attenuated the signaling function of Na/K-ATPase, which subsequently causes inhibition of cell growth. These findings are consistent with previous studies in Na/K-ATPase knockdown cell lines (Liang, Cai et al.
2006, Liang, Tian et al. 2007, Liu, Bai et al. 2012). We also found that ATP1A1-AS1 can be regulated by histone acetylation as well as DNA methylation. We noticed that the change in Na/K-ATPase α1 expression by overexpression of antisense ATP1A1-AS1 transcript is moderate. It was also observed that the expression level of this antisense RNA is much lower than its sense gene. Therefore, the physiological role of ATP1A1-AS1 remains to be elucidated. On the other hand, our result does not exclude the possibility that
Na/K-ATPase α1 may not be the only target of ATP1A1-AS1.
In addition to the findings on a novel endogenous Na/K-ATPase α1 regulator, we also examine the effect of Na/K-ATPase α1 reduction on cardiac function using a CKD animal model. Our results demonstrated that CKD induces cardiac hypertrophy and fibrosis in WT mice. Reduction of Na/K-ATPase α1 significantly reduces cardiac hypertrophy, but
77 it had no significant effect on the formation of cardiac fibrosis. In mechanistic study, we identified that a Na/K-ATPase-related Src/NFB signaling was involved in the regulation of cardiac miR-29b expression in the WT mice, which contributes to the formation of cardiac fibrosis by increasing collagen synthesis. Reduction of Na/K-ATPase α1 blocks the activation of Src and NFB, therefore had no significant effect on the expression of miR-
29b. However, we found that even though Na/K-ATPase reduction in α1+/- mice caused a deficiency in Na/K-ATPase signaling and prevented miR-29b-3p dysregulation, it did not completely block PNx-induced cardiac fibrosis, suggesting that other pathways may contribute to the formation of cardiac fibrosis in this model. Our previous studies showed that cardiac apoptosis increased in α1+/- mice subjected to PNx surgery or MBG infusion
(Liu, Bai et al. 2012, Drummond, Sayed et al. 2014). These apoptotic events may cause the release of cytokines that stimulate fibrotic changes in the tissue. These results also suggest that the reduction of Na/K-ATPase may not be an appropriate strategy to reduce CKD- related cardiac fibrosis.
In summary, our study demonstrated an important role of Na/K-ATPase α1 in maintaining the normal cardiac function. Reduction of Na/K-ATPase α1 could disproportionally block its signaling function and result in cell apoptosis or cell growth inhibition.
78
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