Download Signaling pathways in myocyte hypertrophy. Role of GATA4

SIGNALING PATHWAYS IN
MYOCYTE HYPERTROPHY
RISTO
KE RKE LÄ
Role of GATA4, mitogen-activated protein kinases and
protein kinase C
Department of Pharmacology
and Toxicology and
Biocenter Oulu,
University of Oulu
OULU 2003
RISTO KERKELÄ
SIGNALING PATHWAYS IN
MYOCYTE HYPERTROPHY
Role of GATA4, mitogen-activated protein kinases and
protein kinase C
Academic Dissertation to be presented with the assent of
the Faculty of Medicine, University of Oulu, for public
discussion in the Auditorium of the Department of
Pharmacology and Toxicology, on April 11th, at 12 noon.
O U L U N Y L I O P I S TO, O U L U 2 0 0 3
Copyright © 2003
University of Oulu, 2003
Supervised by
Professor Heikki Ruskoaho
Reviewed by
Docent Päivi Koskinen
Docent Veli-Matti Kähäri
ISBN 951-42-6995-0
(URL: http://herkules.oulu.fi/isbn9514269950/)
ALSO AVAILABLE IN PRINTED FORMAT
Acta Univ. Oul. D 719, 2003
ISBN 951-42-6994-2
ISSN 0355-3221
(URL: http://herkules.oulu.fi/issn03553221/)
OULU UNIVERSITY PRESS
OULU 2003
Kerkelä, Risto, Signaling pathways in myocyte hypertrophy. Role of GATA4, mitogenactivated protein kinases and protein kinase C
Department of Pharmacology and Toxicology and Biocenter Oulu, University of Oulu, P.O.Box
5000, FIN-90014 University of Oulu, Finland
Oulu, Finland
2003
Abstract
Cardiac myocytes react to increased workload and hypertrophic neurohumoral stimuli by increasing
protein synthesis, reinitiating expression of fetal forms of structural genes, α-skeletal actin (α-SkA)
and β-myosin heavy chain (β-MHC), and by increasing expression and secretion of atrial natriuretic
peptide (ANP) and B-type natriuretic peptide (BNP). Initially, the response is beneficial, but when
prolonged, it leads to pathological cardiomyocyte hypertrophy. In this study, cardiomyocyte
hypertrophy was initiated by hypertrophic agonists, endothelin-1 (ET-1) and phenylephrine (PE), and
by increased stretching of atrial wall.
Transcription factor GATA4 was studied to identify the mechanism leading to increased gene
expression of BNP. In BNP promoter, GATA4 binds to cis elements mediating hypertrophic
response. Eliminating GATA4 binding by using the decoy approach, basal BNP gene expression was
reduced. To identify mechanisms regulating GATA4, the roles of mitogen-activated protein kinases
(MAPKs) were studied. Activation of p38 MAPK increased GATA4 binding to BNP gene and led to
increased GATA4 dependent BNP gene expression. p38 MAPK was required for ET-1 induced
GATA4 binding, whereas extracellular signal-regulated kinase (ERK) was required for maintaining
basal GATA4 binding activity. PE and ET-1 activated protein kinase C (PKC) signaling in cardiac
myocytes. Antisense oligonucleotide inhibition of PKCα markedly reduced PE induced ANP
secretion and ET-1 induced BNP secretion, whereas gene expression of natriuretic peptides was not
affected. Antisense PKCα treatment inhibited PE induced expression of α-SkA, while increased
protein synthesis or β-MHC gene expression were not affected. Sretching of the perfused rat atria
increased BNP, c-fos and BNP gene expression via mechanism involving p38 MAP kinase activation
of transcription factor Elk-1. In cultured neonatal rat atrial myocytes stretch induced BNP gene
expression was dependent upon transcription factor Elk-1 binding sites within the BNP gene
promoter.
In conclusion, hypertrophic signaling in cardiac myocytes involves multiple signaling cascades.
Activation of p38 MAPK is required for the development of ET-1 induced hypertrophic phenotype
and GATA4 mediated BNP gene expression in cultured ventricular myocytes, and for stretch induced
Elk-1 dependent BNP gene expression in atrial myocytes. PKCα is involved in PE induced
hypertrophic response and PE induced switch in gene programming inducing expression of α-SkA,
the fetal form of cardiac α-actin.
Keywords: endothelin-1, GATA4, hypertrophy, mitogen-activated protein kinase, protein
kinase C
To my family
Acknowledgements
This work was carried out during the years 1996–2003 at the department of
Pharmacology and Toxicology in the University of Oulu. Professor Heikki Ruskoaho, my
supervisor in this research work, has been an endless source of inspiration. Having time
and patience to listen and to guide in every minor issue, it has also been refreshing to
share ideas and experiences on fishing and ice hockey with him. The relaxing atmosphere
among the group members has helped me carry out the thesis work in his group.
I wish to thank Professor Olavi Pelkonen, the head of the Department of
Pharmacology and Toxicology, whose way of leading the Department has provided an
encouraging athmosphere and excellent conditions for scientific work. I also appreciate
the important collaboration with Professors Juhani Leppäluoto, Olli Vuolteenaho and
Matti Weckström. Their methodological advice and instructive comments have been
essential for the completion of my work.
I am grateful to Docents Veli-Matti Kähäri and Päivi Koskinen for the careful review
of this thesis. I express my thanks to Anna Vuolteenaho for revising the language of this
thesis.
My warmest thanks are due to Dr. Heikki Tokola and Dr. Marja Luodonpää for
introducing me to the wonderful world of neonatal rat ventricular cell culture. I also want
to express my gratitude to Docent Olli Arjamaa for patient and detailed introduction to
isolated rat perfused atrial preparations. Special thanks to Dr. Sampsa Pikkarainen for the
hilarious moments during, between and after the overwhelming number of experiments.
It has truly been a pleasure to work with you.
I want to thank all my co-authors and collaborators: Sari Bergström, Mika Ilves,
Theresa Majalahti-Palviainen, Juhani Pöntinen and Jarkko Ronkainen. I also wish to
thank Sinikka Eskelinen and Juha Tuukkanen for valuable advice in microscopy imaging.
The professional work by Marja Arbelius, Kati Viitala, Sirpa Rutanen and Anneli
Rautio has been most valuable and essential for the completion of the projects. I also own
my warmest gratitude to Helka Koisti, Pirjo Korpi, Tuulikki Kärnä, Tuula Lumijärvi,
Kaisa Penttilä and Ulla Weckström for outstanding technical assistance. Raija Hanni, Esa
Kerttula and Kauno Nikkilä have been of most valuable help in issues dealing with
everyday work in the Department. I also express my gratitude to Terttu Keränen and
Marja Räinä from the instrument care unit, as well as Tuula Inkala and the rest of the
personnel at the Center of Experimental Animals. The skilled expertise and friendly cooperation of Liisa Kärki and Seija Leskelä from the photography lab also deserves my
thanks.
I sincerely thank Dr. Jarkko Piuhola for his friendship, and look forward to pursuing
matching his fast reflexes both in the lab and on the track. I also want to thank Jani Aro,
Gábor Földes, Jukka Hakkola, Nina Hautala, Pietari Kinnunen, Zoltán Lakó-Futó, Hanna
Leskinen, Jarkko Magga, Minna Marttila, Antti Ola, Tuomas Peltonen, Chris Pemberton,
Tanja Rauma, Hannu Romppanen, Jaana Rysä, Balázs Sármán, István Szokodi, Réka
Skoumal, Maria Suo, Olli Tenhunen and all my colleagues in the Department of
Physiology for nice conversations and creating and supporting the most valuable team
spirit.
I want to thank all my friends for support during the course of this study. Special
thanks to Pekka Puolakka for providing an ancillary hand in the cell culture.
My parents Arja and Aki Kerkelä, as well as my brother Tuomo, are warmly
ackownledged for encouragement at all times. I want to thank my grandparents Saima
and Eero Kerkelä for support and constant interest in my work. I also wish to thank
Kyllikki and Erkki Riihimäki for their care and the relaxing moments during holidays.
Finally, and most of all, I want to express my deepest gratitude to my beloved wife
Seija for her love and support.
This work was supported financially by Biocenter Oulu, the Academy of Finland,
Sigrid Juselius Foundation, the Foundation of Oulu University, Aarne Koskelo
Foundation, the Research Foundation of Orion, the Research and Science Foundation of
Farmos, Paulo Foundation, Pharmacal Research Foundation, Finland, Duodecim Society
of Oulu, Ida Montin Foundation, Finnish Cultural Foundation and the Finnish Heart
Foundation.
Oulu, March 2003
Risto Kerkelä
Abbreviations
AM
Ang II
ANP
AP-1
α-SkA
ATF-2
β-MHC
BNP
cAMP
cDNA
cGMP
CHF
CMV
CNP
CsA
EBS
ERK
ET
ETx
FOG
GAPDH
G-proteins
GPCR
GSK3
GTP
HIF-1
iNOS
IL-1
IP3
IR
JNK
adrenomedullin
angiotensin II
atrial natriuretic peptide
activator protein-1
skeletal α-actin
activating transcription factor-2
β-myosin heavy chain
B-type natriuretic peptide
adenosine 3´,5´-cyclic monophosphate
complementary deoxyribonucleic acid
cyclic guanosine monophosphate
congestive heart failure
cytomegalovirus
C-type natriuretic peptide
cyclosporin A
ETS binding sequence
extracellular signal-regulated kinase
endothelin
endothelin receptor subtype
friend of GATA
glyceraldehyde 3-phosphate dehydrogenase
guanine nucleotide binding proteins
G-protein coupled receptor
glycogen synthase-3 kinase
guanosine triphosphate
hypoxia-inducible factor-1
inducible nitric oxide synthase
interleukin-1
inositol-1,4,5-triphosphate
immunoreactive
c-Jun N-terminal kinase
LV
LVH
MAPK
MAPKK
MAPKKK
MBP
MEF2
MHC
MI
MKP
MLC
mRNA
NFAT
NEP
NPR
PCR
PE
PKC
PMA
PI3K
RAS
RIA
RT-PCR
SAPK
SD
SEM
SRE
SRF
STAT
TGF-β
TnC
TNF-α
TnI
TnT
left ventricular
left ventricular hypertrophy
mitogen-activated protein kinase
mitogen-activated protein kinase kinase
mitogen-activated protein kinase kinase kinase
myelin basic protein
myocyte-specific enhancer-binding factor-2
myosin heavy chain
myocardial infarction
mitogen-activated protein kinase phosphatase
myosin light chain
messenger ribonucleic acid
nuclear factor of activated T-cells
neutral endopeptidase
natriuretic peptide receptor
polymerase chain reaction
phenylephrine
protein kinase C
phorbol myristate acetate
phosphatidylinositol-3-OH kinase
renin-angiotensin system
radioimmunoassay
reverse transcriptase polymerase chain reaction
stress-activated protein kinase
Sprague-Dawley
standard error of mean
serum response element
serum response factor
signal transducers and activators of transcription
transforming growth factor-beta
troponin C
tumor necrosis factor-alpha
troponin I
troponin T
α
β
γ
δ
ε
η
ι
θ
ζ
λ
alpha
beta
gamma
delta
epsilon
eta
iota
theta
zeta
lambda
List of original papers
This thesis is based on the following articles, which are referred to in the text by Roman
numerals:
I
Pikkarainen, S, Kerkelä, R, Pöntinen, J, Majalahti-Palviainen, T, Tokola, H,
Eskelinen, S, Vuolteenaho, O & Ruskoaho, H (2002) Decoy oligonucleotide
characterization of GATA4 transcription factor in hypertrophic agonist induced
responses of cardiac myocytes. J Mol Med. 80: 51–60.
II
Kerkelä, R, Pikkarainen, S, Majalahti-Palviainen, T, Tokola, H & Ruskoaho, H
(2002) Distinct Roles of Mitogen Activated Protein Kinase Pathways in GATA4
Transcription Factor Mediated Regulation of B-type Natriuretic Peptide Gene. J Biol
Chem. 277: 13752–60.
III
Kerkelä, R, Ilves, M, Pikkarainen, S, Tokola, H, Ronkainen J, Vuolteenaho, O,
Leppäluoto, J & Ruskoaho, H (2002) Identification of PKC alpha Isoform Specific
Effects in Cardiac Myocytes Using Antisense Phosphorothioate Oligonucleotides.
Mol Pharmacol. 62: 1–10.
IV
Kerkelä, R, Bergström S, Ilves, M, Vuolteenaho, O, Leppäluoto, J & Ruskoaho, H
MAPK regulation of cardiac gene expression in response to atrial stretch. Manuscript.
Contents
Abstract
Acknowledgements
Abbreviations
List of original papers
1 Introduction ...................................................................................................................15
2 Review of literature .......................................................................................................17
2.1 The genetic response of heart to cardiac overload..................................................17
2.1.1 Natriuretic peptides..........................................................................................18
2.1.2 Sarcomeric proteins .........................................................................................21
2.2 Signaling pathways in cardiac hypertrophy............................................................22
2.2.1 Small GTP-binding proteins ............................................................................22
2.2.1.1 Ras family.................................................................................................23
2.2.1.2 Rho family ................................................................................................24
2.2.2 Protein kinase C...............................................................................................24
2.2.2.1 Conventional PKCs ..................................................................................26
2.2.2.2 Novel PKCs ..............................................................................................27
2.2.2.3 Atypical PKCs ..........................................................................................28
2.2.3 Mitogen-activated protein kinases ...................................................................28
2.2.3.1 Extracellular signal-regulated kinase........................................................28
2.2.3.2 c-Jun N-terminal kinase............................................................................30
2.2.3.3 p38 mitogen-activated protein kinase .......................................................31
2.2.4 Phosphatases....................................................................................................33
2.2.4.1 Calcineurin................................................................................................33
2.2.4.2 Mitogen-activated protein kinase phosphatases........................................34
2.3 Cardiac transcription factors...................................................................................34
2.3.1 GATA family of transcription factors ..............................................................35
2.3.2 NFATc family of transcription factors .............................................................36
2.3.3 The ETS-domain transcription factor family ...................................................38
2.3.4 HIF-1 ...............................................................................................................40
3 Aims of the research ......................................................................................................41
4 Materials and methods...................................................................................................42
4.1 Materials.................................................................................................................42
4.2 Experimental animals .............................................................................................43
4.3 Experimental protocols...........................................................................................43
4.3.1 Neonatal rat cardiac myocyte cell culture (I–IV).............................................44
4.3.2 Isolated perfused atrial preparations (IV) ........................................................44
4.4 Isolation and analysis of cytoplasmic RNA (I, III–IV)...........................................45
4.5 Radioimmunoassays (I, III–IV) ..............................................................................46
4.6 Reporter gene assays (I–II, IV)...............................................................................47
4.7 Gel mobility shift assys (I–II, IV)...........................................................................47
4.8 Western blot analysis (II–IV)..................................................................................48
4.9 Kinase activity assays (II, III).................................................................................48
4.10 Calcineurin activity assay (III) .............................................................................49
4.11 Cytochemistry (I, III)............................................................................................49
4.12 Protein synthesis (I–III) ........................................................................................50
4.13 Statistical analysis.................................................................................................50
5 Results ...........................................................................................................................51
5.1 Role of GATA4 in the regulation of BNP gene expression (I)................................51
5.1.1 GATA4 DNA binding activity .........................................................................51
5.1.2 Effect of GATA decoy treatment on ANP and BNP.........................................52
5.1.3 Effect of GATA decoy on protein synthesis and sarcomere assembly .............53
5.2 Role of MAP kinases in GATA4 activation and BNP gene expression (II) ............53
5.2.1 Activation of ERK and p38 MAP kinase by ET-1 ...........................................53
5.2.2 Effect of ERK and p38 inhibition on ET-1 induced protein synthesis .............54
5.2.3 Regulation of GATA4 by MAP kinases ...........................................................54
5.2.4 p38 MAP kinase regulation of GATA-dependent promoter.............................55
5.3 PKCα in the regulation of cardiomyocyte hypertrophy (III) ..................................56
5.3.1 Antisense inhibition of PKCα..........................................................................56
5.3.2 Effect of antisense ODN inhibition of PKCα on hypertrophic phenotype ......57
5.3.3 Effect of antisense PKCα treatment on ET-1 and PE induced natriuretic
peptide secretion and gene expression.............................................................58
5.4 Regulation of atrial gene expression by MAP kinases (IV)....................................61
5.4.1 Effect of MAPK inhibition on stretch induced natriuretic peptide gene
expression and secretion ..................................................................................61
5.4.2 Effect of atrial stretch on cardiac gene expression ..........................................62
5.4.3 Effect of stretch on transcription factors..........................................................62
5.4.4 Regulation of BNP gene expression via EBS ..................................................63
6 Discussion .....................................................................................................................64
6.1 Role of GATA4 in the regulation of myocyte hypertrophy.....................................64
6.2 Distinct roles of MAP kinases in the regulation of myocyte hypertrophy..............65
6.3 PKCα signaling in myocyte hypertrophy ...............................................................67
6.4 MAPK regulation of stretch induced cardiac gene expression ...............................68
6.5 Future perspectives .................................................................................................69
7 Summary and conclusions .............................................................................................70
References
1 Introduction
Primary and acquired structural heart diseases are the leading causes of mortality in the
industrialized world (World Health Organization 2002). Acquired structural heart disease
most often results from myocardial infarction (MI) due to coronary artery disease,
hypertension and hypertensive cardiomyopathy (Berne & Levy 1993). Under stress
compromising cardiac function, the heart attempts to maintain normal contractile
function and cardiac output by undergoing a process of cardiac hypertrophy and
remodeling. Initially, increase in myocyte size and thickening of left ventricle wall is
beneficial, but when prolonged, it leads to pathological hypertrophy (for review, see
Lorell & Carabello 2000). Increased stretching of cardiac myocytes is the main factor
inducing hypertrophic growth, but circulating substances, such as endothelin-1 (ET-1)
and angiotensin II (Ang II) either directly or indirectly also induce hypertrophic growth
of cardiac myocytes (for review, see Sussman et al. 2002). It is not clear whether a
primary stimulus for hypertrophy is the mechanical stretch itself or a concurring increase
in neural and humoral factors. Cardiomyocytes respond to extracellular signals by
hypertrophic growth of individual cells initiated by a specific set of mechanisms that
regulate and modulate gene expression (for review, see Sadoshima & Izumo 1997).
Multiple signaling molecules, many of them same as those that regulate proliferation of
cancer cells or immune cells, are assembled between the signal and the gene to
participate in signaling regulating growth of cardiomyocytes (for review, see Sugden &
Clerk 1998b).
A set of protein kinase cascades has been in the focus of attention in studies on
molecular mechanisms regulating hypertrophic response of cardiac myocytes evoked by
external stimulus. Mechanical stretch of cardiomyocytes has been shown to activate
second messengers, such as protein kinase C (PKC), mitogen-activated protein kinase
cascades (MAPK) and calcineurin, which ultimately affects nuclear factors and the
regulation of a number of genes, including atrial natriuretic peptide (ANP), skeletal αactin (α-SkA) and β-myosin heavy chain (β-MHC) (for review, see Sugden & Clerk
1998b, Molkentin & Dorn II 2001). It is evident that there is no isolated signaling
cascade for each stimulus or response, but the multitude of signaling molecules rather
forms a network of cascades with numerous elements facilitating cross-talk between the
cascades. In this manner it is evident that blockade of any of the intracellular signaling
16
pathways is not sufficient to efficiently prevent the hypertrophic growth of the
cardiomyocyte. Similarly, activation of any of the pathways is predicted to activate
hypertrophic growth of cardiomyocyte by cross-talking and activating other signaling
networks.
The aim of the present study was to evaluate the role of GATA4 transcription factor in
hypertrophic cardiomyocyte growth and gene expression, and to study the significance of
the signaling pathways involved in cardiomyocyte stretch and hypertrophic agonist
induced cardiomyocyte growth. Significance of GATA4 binding to B-type natriuretic
peptide (BNP) gene promoter was studied by using decoy oligonucleotide approach.
Mechanisms leading to GATA4 phosphorylation and activation were further studied by
using specific inhibitors of MAP kinases and plasmids overexpressing MAPK pathway
kinases. Role of PKCα in the regulation of cardiomyocyte hypertrophy was studied by
using antisense oligonucleotides to inhibit PKCα.
2 Review of literature
2.1 The genetic response of heart to cardiac overload
The mammalian myocardium undergoes a period of hypertrophic growth during the
postnatal maturation, which is characterized by an increase in size of individual cardiac
myocytes without cell division (Lorell & Carabello 2000). The pattern of developmental
hypertrophy is reinitiated in the adult heart in response to diverse mechanical,
hemodynamic, hormonal and pathologic stimuli (Hunter & Chien 1999). Increased
mechanical and neurohumoral load, such as hypertension, ischemic heart disease,
valvular insufficiency and cardiomyopathy, increases wall thickness and results in
concentric hypertrophy (Lorell & Carabello 2000). At the cellular level, cardiac myocytes
respond to diverse types of biomechanical stress by initiating several different processes
that via activation of transcription factors lead to hypertrophic gene expression and
growth of individual myocytes (Fig. 1). Initially, the response is beneficial, but when
prolonged, it leads to pathological myocyte hypertrophy. The first genetic response to
increased load is activation of a pattern of early response, or immediate early genes: cfos, c-myc and c-jun (Yamazaki et al. 1995b; Table.1). This is followed by induction of
certain genes, such as ANP, BNP, β-MHC and α-SkA, accompanied by development of
hypertrophic phenotype characterized by an increase in cell surface area, protein
concentration and protein to DNA ratio (Sadoshima & Izumo 1997).
18
Fig. 1. A simplified model of a signal transduction pathway initiated by mechanical stretch or
neurohumoral agonists in cardiac myocytes. The arrows provide the directional information
of the signals forming a hierarchic system between the signaling molecules inside the cell.
GPCR, G-protein-coupled receptor; PE, phenylephrine; MAPKKK, mitogen-activated
protein kinase kinase; NFATc4, nuclear factor of activated T-cells c4; FOG-2, friend of
GATA-2
2.1.1 Natriuretic peptides
ANP, BNP and C-type natriuretic peptide (CNP) are the known members of the
mammalian natriuretic peptide system. Studies concerning ANP go back to 1981, when it
was found that intravenous administration of atrial extracts into intact rats causes diuresis
and natriuresis (de Bold et al. 1981). Soon thereafter, ANP molecule was purified and
sequenced (Atlas et al. 1984). A few years later BNP and CNP were discovered from the
porcine brain (Sudoh et al. 1988, Sudoh et al. 1990). ANP and BNP are cardiac
hormones, and involved in the regulation of blood pressure and fluid homeostasis (Nakao
19
et al. 1992a). CNP is mainly found in the central nervous system and vascular endothelial
cells (Chen & Burnett, Jr. 1998), where it increases cGMP levels and causes
vasorelaxation (Furuya et al. 1990). Natriuretic peptides share a common structural motif,
consisting of a 17-amino acid loop formed by an intramolecular disulphide linkage
between two cysteine residues (for review, see Yandle 1994). The biologically active
human ANP shares a high homology with other species, as does the active CNP. In
contrast, the sequence variability of BNP between species is large, and the predominant
circulating forms of BNP are 26, 45 and 23 amino acid peptides in pigs, rats and humans,
respectively (Nakao et al. 1992a).
ANP and BNP are present as single-copy genes and are organized into three exons and
two introns (Greenberg et al. 1984, Seidman et al. 1984, Argentin et al. 1985, Seilhamer
et al. 1989). The ANPs are derived from a common precursor, named preproANP, which
contains 149–152 amino acids depending on the species. Each preproANP molecule
contains a signal peptide sequence at its amino terminal end. ANP is stored in atrial
secretory granules mainly as a 126-amino acid peptide, proANP1-126, which is formed by
cleavage of the signal peptide (Thibault et al. 1987). Upon release of ANP1-126 from
secretory granules, the hormone is further split into an amino-terminal fragment,
proANP1-98, and the biologically active carboxy-terminal peptide, ANP99-126 (or ANP1-28)
(Schwartz et al. 1985, Thibault et al. 1985). The mRNA of BNP and CNP are also
translated to preproBNP and preproCNP (Nakao et al. 1992a), which are further reduced
by the cleavage of the signal peptide (for review, see Nakao et al. 1992a). The major
storage form of BNP in the heart is cleaved mature peptide (Saito et al. 1989,
Kambayashi et al. 1990, Ogawa et al. 1990, Mukoyama et al. 1991).
In the normal adult heart, ANP is mainly synthesized in the atria, but small quantities
of ANP mRNA have also been found in normal adult ventricular myocytes, where the
levels are approximately 0.5 to 3% of that in the atria (Ruskoaho 1992). Expression of
atrial ANP is initially low and increases through development. In contrast, ventricular
myocytes from neonatal hearts contain substantial amounts of ANP mRNA which
declines rapidly after birth (Ruskoaho 1992). The ventricular expression and secretion of
ANP is substantially elevated in cardiac hypertrophy. The major stimulus regulating ANP
release is myocyte stretch (Lang et al. 1985, Ruskoaho et al. 1986). The secretion of ANP
from the granules is also induced by a number of hormones and vasoactive peptides (for
review, see Ruskoaho 1992).
BNP is expressed in both atria and ventricles, but is mainly released from the
ventricles (Hosoda et al. 1991, Ogawa et al. 1991). BNP transcripts have also be detected
in the central nervous system, lung, thyroid, adrenal gland, kidney, spleen, small
intestine, ovary, uterus, and striated muscle (Gerbes et al. 1994). In studies with rat and
man, the distribution of BNP has been strong in the spinal cord, while only small amounts
are detected in the brain (Aburaya et al. 1989, Aburaya et al. 1991, Nakao et al. 1992a).
In all peripheral tissues, the level of both natriuretic peptide transcripts are approximately
1–2 orders of magnitude lower than in cardiac ventricular tissues (Gerbes et al. 1994).
Ventricular levels of BNP are substantially increased in response to chronic cardiac
overload in the human heart (Hosoda et al. 1991, Takahashi et al. 1992).
CNP is a 22-amino-acid peptide, structurally related to, but genetically distinct from
ANP and BNP (Minamino et al. 1991, Komatsu et al. 1991). Although initially identified
in porcine brain, CNP immunoreactivity has been found in human vascular endothelial
20
cells, plasma, and kidney (Sudoh et al. 1990, Chen & Burnett, Jr. 1998). CNP has potent
systemic cardiovascular actions, including reductions in cardiac filling pressures and
output secondary to vasorelaxation and decreases in venous return, but has minimal renal
actions (for reviews, see Needleman et al. 1989, Rosenzweig & Seidman 1991, Ruskoaho
1992, Yandle 1994, Nakao et al. 1996, Chen & Burnett, Jr. 1998). Unlike ANP, CNP has a
direct biological role as a regulator of cardiovascular tone (Wei et al. 1993).
Three natriuretic peptide receptors (NPRs) have been described that have different
binding capacities for ANP, BNP and CNP (for reviews, see Espiner et al. 1986,
Ruskoaho 1992, Anand-Srivastava 1997, Levin et al. 1998). Two of the receptors, NPRA
(CGA) and NPRB (CGB), are guanylyl cyclases–linked receptors, and mediate the biologic
actions, whereas NPRC is not coupled to guanylyl cyclase (Chang et al. 1989, Chinkers et
al. 1989, Schulz et al. 1989, Jamison et al. 1992). The natural ligands of NPRA are ANP
and BNP, whereas that of NPRB is CNP (Jamison et al. 1992). The rank order of ligand
selectivity for NPRA receptor is ANP ≥ BNP > CNP, and for NPRB receptor
CNP > ANP ≥ BNP (Nakao et al. 1992b). The third receptor, NPRC, binds all three
natriuretic peptides. Its messenger RNA lacks the guanylyl cyclase sequence present in
the mRNA of the other natriuretic peptide receptors, suggesting that the principal
function of NPRC is to remove natriuretic peptides from the circulation (for review, see
Anand-Srivastava & Trachte 1993). Natriuretic peptides, CNP in particular, are also
subject to degradation by the ectoenzyme neutral endopeptidase 24.11 (NEP), which is
widely distributed in the kidney, lung, heart, and endothelial cells (Kenny et al. 1993,
Brandt et al. 1997). Natriuretic peptide receptor binding sites have been localized to atrial
and ventricular endocardium, as well as aorta, pulmonary arteries and epicardial
mesothelium (Rutherford et al. 1992). The significance of natriuretic peptide receptors
has been mainly revealed in studies with transgenic mice. NPRA deficient mice display an
elevated blood pressure resulting in severe cardiac hypertrophy, whereas introduction of
the NPRA transgene in the cardiac myocytes reduced cardiac myocyte size in both wildtype and originally NPRA null mice (Kishimoto et al. 2001).
Table 1. Changes in gene expression associated with cardiac hypertrophy and congestive
heart failure (Sadoshima & Izumo 1997, Lorell & Carabello 2000, Wilson & Spinale
2001, Sjaastad et al. 2003). AM, adrenomedullin; CT-1, cardiotrophin-1, NCX, Na+-Ca2+
exchanger; MMP, matrix metalloproteinase, SERCA, sarco(endo)plasmic reticulum Ca2+
ATPase.
Early response
c-fos
c-myc
c-jun
BNP
Egr-1
Intermediate response
ANP
β-MHC
Skeletal α-actin
AM
Phospholamban
Late response
Cardiac α-actin
CT-1
NCX
MMPs
SERCA
Natriuretic peptides have been used in the treatment of diseases, with the most experience
coming from intravenous BNP in the treatment of congestive heart failure (CHF)
(McMurray & Pfeffer 2002, Mills et al. 2002). Another pharmacological approach being
used is the inhibition of natriuretic peptide metabolism by NEP inhibitor drugs, which are
currently being investigated as treatment for CHF and systemic hypertension (Corti et al.
21
2001, Weber 2001, McMurray & Pfeffer 2002). Increased blood levels of natriuretic
peptides, ANP and BNP, have been found in certain disease states, suggesting a role in
the pathophysiology of those diseases, including CHF, systemic hypertension, and acute
MI (Ruskoaho 1992, Swynghedauw 1999). Elevated levels of ANP have been identified
as indicators of adverse prognosis in the setting of CHF and MI (Rossi et al. 2000). BNP
has proved to be a useful tool in the diagnosis of heart failure in the community and in the
detection of asymptomatic left ventricular (LV) dysfunction (Daly et al. 2002). In
addition, BNP plasma level values have been suggested as of guidance tool for
pharmacotherapy in heart failure (Nicholls et al. 2001).
Studies with different in vivo and in vitro models have revealed that BNP gene
expression is regulated by transcriptional and post-transcriptional mechanisms (Tokola et
al. 2001, Suo et al. 2002). Transcriptional regulation of the BNP gene involves binding of
transcription factors, such as GATA4, AP-1 (activator protein-1), transcription enhancer
factor-1, Nkx-2.5 and NFATc4, to the 5’-flanking sequence of the BNP gene promoter
(Grepin et al. 1994, Thuerauf et al. 1994, Thuerauf & Glembotski 1997, Durocher et al.
1997). The signaling cascades responsible for activation of transcription factors are not
fully elucidated, but they involve activation of intracellular protein kinases and
phosphatases, such as PKC, MAPKs and calcineurin, as discussed later in detail (for
reviews, see Sugden & Clerk 1998b, Molkentin & Dorn II 2001).
2.1.2 Sarcomeric proteins
Increased ventricular load requires enhanced cardiac myocyte contractile performance,
which is preceded by an increase in force-generating units (sarcomeres) in the myocyte
(Lorell & Carabello 2000). The sarcomere consists of thick and thin filaments, which are
organized to form contractile units (Murphy 1996). The thick filament proteins are two
myosin heavy chain proteins (α-MHC and β-MHC), two myosin light chain proteins
(MLC-1 and MLC-2) and C-protein (Hoh et al. 1978, Whalen & Sell 1980, Kasahara et
al. 1994). β-MHC is the dominant isoform of the heavy chains in the ventricles of
humans throughout the development (Schier & Adelstein 1982, Wessels et al. 1991,
Mercadier et al. 1983). In response to mechanical load, genes from various striated
muscle types respond by inhibiting expression of α-MHC, and increasing expression of βMHC (Swynghedauw 1999). In experimental aortic stenosis model increased levels of βMHC mRNA are observed two days after operation, peaking at days 4 to 6 (Izumo et al.
1987). β-MHC gene expression is also elevated in cultured rat cardiac myocytes by
mechanical stress and by neurohumoral activators, such as ET-1 and α1-adrenergic
agonists (Waspe et al. 1990, Shyu et al. 1995, Finn et al. 2001). Studies with rat β-MHC
promoter have revealed that β-MHC gene can be induced by various agonists, including
constitutively active PKC-β isozyme and activated MAPKs (Kariya et al. 1991, Finn et
al. 2001). During embryogenesis, the β-MHC gene is expressed as part of the cardiac
myogenic program under the control of NKX-2.5, myocyte-specific enhancer-binding
factor-2C (MEF2C), and GATA4 (Hasegawa et al. 1997, Skerjanc et al. 1998). There is
also evidence that calcineurin and NFATc4 are involved in the regulation of β-MHC gene
22
expression (Delling et al. 2000). MLC-2 mRNA levels are also upregulated in response to
hypertrophic stimuli by α1-adrenergic agonist PE, phorbol myristate acetate (PMA) and
insulin growth factor-1 (Lee et al. 1988a, Shubeita et al. 1992, Ito et al. 1993).
The thin filament proteins include α- and β-tropomyosin genes, slow-twitch skeletal
and cardiac troponin I genes, troponin C, slow-twitch skeletal and cardiac troponin T
genes and three α-actin genes; α-cardiac, α-skeletal and smooth muscle α-actin, of which
the α-cardiac actin predominates in the adult heart (Cummins & Perry 1973, Toyota &
Shimada 1981, Mayer et al. 1984, Carrier et al. 1992, Jin et al. 1992, Murphy 1996).
Changes in myocardial contractility and relaxation appear to involve changes in troponin
isoform expression and phosphorylation (Solaro & Rarick 1998). The skeletal mRNA
isoform of α-actin (α-SkA) has been found transiently expressed during pressure overload
in rats (Schwartz et al. 1986, Izumo et al. 1988). The activation of vascular smooth
muscle α-actin has also been suggested to occur during aortic constriction induced
cardiac hypertrophy (Black et al. 1991). Levels of α-SkA are elevated at day 2 and reach
peak levels 4 days after the onset of coarctation (Izumo et al. 1988). Hypertrophic
agonists, such as PE and ET-1, have been shown to induce α-SkA gene expression via
pathways involving MAPKs, calcineurin and PKC (Ito et al. 1991, Komuro et al. 1991,
Fuller et al. 1997, Bueno et al. 2002b).
2.2 Signaling pathways in cardiac hypertrophy
The hypertrophic phenotype of cardiomyocytes is initiated by multiple endocrine,
autocrine and paracrine factors that stimulate receptors bound to cell membrane. The
activation of receptors affects multiple cytoplasmic signaling pathways, which regulate
the activity of transcription factors, and finally regulate gene expression. The major
contribution to studying the signaling pathways activated in cardiac hypertrophy was the
development of a cell culture model with cardiac myocytes from neonatal rat hearts
(Simpson & Savion 1982, Simpson et al. 1982). Initially, α1-adrenergic agonists were
shown to induce hypertrophic growth of a myocyte (Simpson 1985). Later, mechanical
stress and several autocrine/paracrine growth factors and neurotransmitters, such as ET-1
and Ang II, were shown to promote a hypertrophic phenotype (Shubeita et al. 1990). For
reviews, see Sugden & Clerk 1998b, Molkentin & Dorn II 2001. There is evidence that
the action of these ligands on cultured neonatal cardiomyocytes mimics cardiac
hypertrophy seen in vivo. During recent years development of kinase inhibitors and usage
of adenoviral vectors and different transgenic models, have made it feasible to study
cardiac load induced hypertrophic signaling in vivo as well.
2.2.1 Small GTP-binding proteins
Small GTP-binding proteins provide a critical link between receptors at the cell surface
and kinase cascades that regulate a variety of cellular processes (Fig. 2). Studies have
identified five different subfamilies: Ras, Rho, ARF, Rab and Ran (Sugden & Clerk
23
2000). When these proteins are GTP-ligated, they are in active state, and when GDPligated, they return to inactive state. To date, participation of members of Ras and Rho
families of small GTP-binding proteins has been described in hypertrophic signaling in
cardiac myocytes (Sugden & Clerk 2000, Nicol et al. 2000).
GPCR agonists
Cardiac contractility Protein
Cell size
synthesis
Hypertrophic
gene expression
Cell
Morphology
Fig. 2. Schematic presentation of myocyte responses by small GTP-binding proteins Ras,
RhoA and Rac1 (modified from Clerk & Sugden 2000).
2.2.1.1 Ras family
Ras is a low molecular weight (21 kDa) guanine nucleotide-binding protein, which is
activated by GDP to GTP exchange initiated by membrane bound receptors. In cardiac
myocytes GTP loading of Ras proteins can be induced by a number of hypertrophic
agonists, such as PE, ET-1 and phorbol esters (Sugden & Clerk 2000). Activation of Ras
can promote activation of Raf-1, phosphatidylinositol-3-OH kinase (PI3K) and Ral.GDS
(Vojtek & Der 1998, Sugden & Clerk 1998b, Molkentin & Dorn II 2001). Ras has been
shown to primarily promote ERK activation, but it can also activate c-Jun N-terminal
kinase (JNK) and p38 MAPKs indirectly through Rac/Cdc42 (Sugden & Clerk 1998b).
Transfection of constitutively activated Ras into cardiac myocytes is sufficient to
promote the hypertrophic phenotype, including increased c-fos, α-SkA and ANP gene
expression and increased sarcomeric organization (Thorburn et al. 1993, Hoshijima et al.
1998, Fuller et al. 1998). In contrast, dominant negative Ras can block PE induced
activation of ERK and ANP gene expression (Thorburn 1994). In vivo cardiospecific
activation of activated Ras in mice leads to development of cardiac hypertrophy (Hunter
24
et al. 1995). Some studies suggest that Ras is a global regulator of gene expression rather
than an activator of the specific hypertrophic program (Sugden & Clerk 2000).
2.2.1.2 Rho family
Members of the Rho family of small GTP-binding proteins are Rac, RhoA and Cdc42
(Sugden & Clerk 1998b). Rac1 can be activated by active Ras, which stimulates PI3K
and increases levels of phosphatidylinositol -1,4,5-triphosphate leading to activation of
Rac1 (Sugden & Clerk 2000). Another mechanism involved is Ras induced activation of
p120GAP and p190GAP leading to Rho activation (Vojtek & Der 1998). RhoA and Rac
have both been implicated in the hypertrophic growth response, mediating both
morphological changes and the changes in gene expression (Clerk et al. 2001).
Expression of activated Rac1 has been shown to stimulate the hypertrophic program,
while expression of dominant negative Rac1 has been inhibitory (Aikawa et al. 1999,
Pracyk et al. 1998). Rac1 has been shown to stimulate the ERK cascade either by
promoting the phosphorylation of c-Raf or by increasing MEK1/2 association with c-Raf
to facilitate MEK1/2 activation (Clerk et al. 2001). Rac1 may also promote JNK
activation, whereas activation of p38 MAPK by Rac1 has not been observed.
A role for RhoA in myocyte hypertrophy is supported by several studies demonstrating
the effects of activated and dominant negative forms of RhoA on hypertrophic gene
expression (Aikawa et al. 1999, Hoshijima et al. 1998, Sah et al. 1996, Thorburn et al.
1997). Interestingly, cardiac specific expression of constitutively active RhoA was not
sufficient to induce hypertrophic phenotype, but it did induce atrial enlargement
associated with decreased heart rate and sinus and atrioventricular nodal dysfunction (Sah
et al. 1999). The effects of RhoA on the morphological aspects of hypertrophy are not
clear, since the results are contradictory (Thorburn et al. 1997, Hoshijima et al. 1998).
RhoA does not stimulate JNK or p38 activity, but it may regulate gene expression by a
mechanism involving Rho-dependent kinase ROK (Nicol et al. 2000, Clerk et al. 2001).
2.2.2 Protein kinase C
Protein kinase C represents a family of at least 12 serine/threonine kinases that participate
in signal transduction in response to multiple stimuli (Nishizuka 1992). PKC is a single
polypeptide with an N-terminal regulatory region and C-terminal catalytic region (Hug &
Sarre 1993, Mellor & Parker 1998). Different isoforms can be divided into three groups
based on differences in their structure, substrate specificity and calcium responsiveness:
conventional or classic (cPKCs; α, β1/2 and γ), novel (nPKCs; δ, ε, η and θ) and atypical
PKCs (aPKCs; ζ and λ/ι) (Fig. 3). PKC isoforms are widely distributed in mammalian
tissues and some isoforms are restricted to specific tissues. For example, PKCγ is
restricted mainly to the central nervous system and spinal cord, and PKCθ to skeletal
muscle and hematopoietic cells (Nishizuka 1992, Kanashiro & Khalil 1998). Cardiac
25
myocytes express members of all three subfamilies of PKC: one classic isoform (PKCα),
two novel isoforms (PKCδ and PKCε), two atypical isoforms (PKCζ and PKCλ/ι) and
probably PKCβI and PKCβII (Disatnik et al. 1994, Rybin et al. 1997, Clerk & Sugden
2001, Mackay & Mochly-Rosen 2001). Within a single cell, there are also differences in
distribution between PKC isoforms before and after their activation. For example, in rat
cardiac myocytes activated PKCα has been shown to translocate from the cytosolic
fraction to the perinuclear region, whereas PKCε is translocated from the nucleus to
myofibrils (Disatnik et al. 1994). Activation and translocation of the PKC isoforms to
distinct subcellular sites is believed to confer specific physiological actions for each PKC
isoform. The translocation of PKC isozymes involves binding to specific anchoring
proteins, named RACKs (receptors for activated C kinases) (Disatnik et al. 1994,
Mochly-Rosen 1995). Binding of PKC to RACK is saturable and isoform specific, and
within a cell there may be multiple RACKs for each PKC in multiple cellular
compartments (Schechtman & Mochly-Rosen 2001).
V1
C1
C2
Kinase
V5
cPKC (α, β, γ)
nPKC (δ, θ)
aPKC (λ/ι, ζ)
Regulatory
domain
Catalytic
domain
Fig. 3. Domain structure of the PKC subfamilies (modified from Mellor et al. 1998). The
figure shows a comparison of the protein architecture of the various subgroups of the PKC
subfamily, all consisting of constant (C) and variable (V) regions.
26
2.2.2.1 Conventional PKCs
Conventional isoforms of PKC are Ca2+-dependent and activated by both
phosphatidylserine and the second messenger diacylglycerol (Nishizuka 1992, Mellor &
Parker 1998). Phorbol esters such as PMA are widely used as substitutes for
diacylglycerol since they are not rapidly metabolized and produce a robust and long
lasting activation of cPKCs and nPKCs. In the heart, PMA rapidly and irreversibly
translocates cPKCs from the cytosolic fraction to cell membrane (Rybin & Steinberg
1994, Clerk et al. 1995). Both ET-1 and PE stimulate the hydrolysis of the membrane
phospholipid, phosphatidylinositol 4,5-bisphosphate, leading to formation of IP3 and
diacylglycerol, the physiological activator of cPKCs and nPKCs (Steinberg et al. 1989,
Hilal-Dandan et al. 1992).
PKCα is the major calcium-dependent PKC isozyme present in neonatal cardiac
myocytes. In resting cells PKCα is located in soluble fraction, and increase in calcium
concentration translocates PKCα to the particulate fraction (Goldberg et al. 1997, Muth et
al. 2001). Exposure of cardiac myocytes to PMA or α-adrenergic receptor agonist
norepinephrine (NE) has been shown to translocate PKCα to the perinuclear membrane
(Disatnik et al. 1994). In contrast, translocation of PKCα is not detected following
exposure of cardiac myocytes to ET-1 or PE (Clerk et al. 1994). PKCβ1 is located in the
cytoplasm and around the nucleus in resting cells, but activation with PMA translocates
PKCβ1 into nucleus (Disatnik et al. 1994). Activated PKCβ2 translocates from
cytoskeletal elements to perinuclear region and cell periphery (Disatnik et al. 1994).
PKCα is detected in extracts from fetal and neonatal rat cardiac myocytes (Rybin &
Steinberg 1994). In contrast, only a small amount of PKCα is detected in the adult rat
hearts and no PKCα immunoreactivity is found in adult ventricular myocytes, suggesting
that in the adult rat heart PKCα is present in nonmyocyte elements (Rybin & Steinberg
1994). However, 2- and 4-month-old transgenic mice overexpressing L-type calcium
channel showed increased membrane association of PKCα (Muth et al. 2001). PKCα can
be constitutively activated by removing the autoinhibitory pseudosubstrate domain and
activating the diacylglycerol binding domain in the N-terminal regulatory region.
Transfection of this constitutively activated mutant of PKCα activates MLC-2 and ANP
gene expression (Muramatsu et al. 1989, Shubeita et al. 1992). Transfection of cardiac
myocytes with PKCβ, which has been similarly constitutively activated, induces β-MHC
gene expression and activates AP-1 dependent transcriptional activity (Kariya et al.
1991). Furthermore, cardiospecific overexpression of PKCβ1 induces activation of the
hypertrophic gene programme (Bowman et al. 1997). On the other hand, studies with
PKCβ1 knockout mice demonstrate that PKCβ1 is not necessary for the development of
cardiac hypertrophy nor does it attenuate the hypertrophic response (Roman et al. 2001).
In transgenic mice PKCβ2 expression under truncated α-MHC promoter results in the
development of hypertrophic phenotype, including increase in myocyte size and
increased expression of β-MHC, c-fos, ANP and TGFβ1 (Wakasaki et al. 1997). Finally, in
samples taken from end-stage human dilated cardiomyopathy, the activity of the classical
isoforms, PKCα and PKCβ1/2, is increased, whereas PKCε activity remains unchanged
27
(Bowling et al. 1999). Studies with PKCβ1 and PKCβ2 translocation inhibitors have been
shown to selectively inhibit phorbol ester induced translocation of PKCβ isozymes, but
not PKCδ or PKCε, demonstrating their potential in studying PKC signaling (Ron et al.
1995).
2.2.2.2 Novel PKCs
Novel isoforms of PKC are Ca2+-independent and activated by both phosphatidylserine
and diacylglycerol. In resting cells, the largest proportion of PKCδ and PKCε is located
in the soluble fraction, and exposure to ET-1 and PE increases their association with the
particulate fraction (Clerk et al. 1994). Activated PKCδ translocates to the perinuclear
region, and PKCε to myofibrils (Disatnik et al. 1994). PKCδ is more abundant in fetal
and neonatal than in adult cardiac myocytes (Rybin & Steinberg 1994). PKCε, which is
present in adult cardiac myocytes, is the major novel isoform associated with
hypertrophic cardiomyocyte growth (Sugden & Clerk 1998b). PKCε has been associated
with cardiac hypertrophy, since PKCε selectively translocates to the particulate fractions
in response to acute and chronic pressure overload and Ang II (Gu & Bishop 1994,
Schunkert et al. 1995, Paul et al. 1997b). PKCδ in turn is activated in vivo by ischemia
and diabetes (Koya & King 1998, Kawamura et al. 1998). Results using the recently
identified PKCδ inhibitor peptide show that cell damage induced by simulated ischemia
is caused, at least in part, by activation of PKCδ (Chen et al. 2001). In neonatal cardiac
myocytes adenoviral overexpression of constitutively active PKCε is sufficient to
increase ANP and β-MHC mRNA levels, but it has no effect on cell size or protein to
DNA ratio (Strait, III et al. 2001). However, adenovirus-mediated expression of dominant
negative PKCε is not sufficient to block ET-1 induced increase in ANP or β-MHC levels
(Strait, III et al. 2001). In transgenic mice expression of PKCε inhibiting peptides results
in increased α-SkA gene expression, increased myocyte size and impaired left ventricular
contractile function (Mochly-Rosen et al. 2000). In contrast, high expression of PKCε
binding peptide ψεRACK, an analogue of the anchoring and activation protein for PKCε,
is associated with increased β-MHC gene expression, decreased myocyte cell size and
normal left ventricular function (Mochly-Rosen et al. 2000). PKCε has also been shown
to regulate activity of other signaling pathways, such as ERK, providing an example of
cross-talk between the signaling molecules (Strait, III et al. 2001). Stress related factors,
such as UV irradiation, ischemia and oxidants have been observed to activate PKCε and
stress activated MAPKs, JNK and p38 MAPK, in parallel (Chen et al. 1999, Brodie et al.
1999, Ping et al. 1999). Taken together, activation and downstream signaling via the
PKCε and PKCδ cascade may be necessary for the induction of hypertrophic gene
programme.
28
2.2.2.3 Atypical PKCs
Atypical isoforms of PKC are Ca2+-independent and do not require diacylglycerol for
activation, yet phosphatidylserine regulates their activity. PKCζ, the atypical isoform
present in cardiac myocytes, is located around the cytosol and the nucleus, and exposure
of cells to PMA translocates PKCζ to perinuclear region (Disatnik et al. 1994). No
translocation of PKCζ is detected following exposure to PE or ET-1 (Clerk et al. 1994).
PKCζ is most abundant in fetal and neonatal myocytes, whereas in the adult rat heart
PKCζ resides in nonmyocyte elements (Rybin & Steinberg 1994). PKCζ has also been
connected with cardiac hypertrophy, since in guinea-pigs aortic banding increases
expression of PKCζ isozyme, and overexpression of constitutively active PKZζ is
sufficient to induce ANP promoter activity in cultured cardiac myocytes (Decock et al.
1994, Jalili et al. 1999).
2.2.3 Mitogen-activated protein kinases
There are multiple MAPK pathways in all eukaryotic cells, which allow the cells to
respond differently to divergent inputs. MAPK signaling cascades are usually divided
into three parallel pathways: ERK, JNK and p38 MAPK pathways. All MAPK pathways
include three signaling levels, i.e. MAPKKK activating MAPKK, which in turn activates
MAPK. Activation of MAPKKKs results from translocation, oligomerization and
phophorylation by upstream kinases (Marshall 1995, Luo et al. 1996, King et al. 1998).
Active MAPKKKs phosphorylate serine and threonine residues in MAPKKs, which in
turn activate tyrosine and threonine residues in the activation loop of the MAPKs. Most
physiological substrates of the MAPKs possess specific binding sites for MAPKs that
allow strong interactions with selectivity for MAPK subfamilies (Kallunki et al. 1996,
Yang et al. 1998a, Deacon & Blank 1999, Tanoue et al. 2000). MAPKs, in turn, also
possess complementary docking sites, which allow them to interact with MAPK binding
domains on substrate proteins (Kallunki et al. 1994, Tanoue et al. 2000). The signaling
mechanism is coordinated by the interaction of components of the protein kinase cascade
with scaffold proteins, such as JNK interacting protein-1 and MEK1 partner (Dickens et
al. 1997, Teis et al. 2002)
2.2.3.1 Extracellular signal-regulated kinase
The ERK family of MAPKs (ERK1,2), also known as p42/44 MAP kinase, is activated
by mitogenic stimuli and generally associated with cell growth and survival. It was the
first of the MAPK pathways proposed to participate in hypertrophic signaling in cardiac
myocytes (Bogoyevitch et al. 1993, Sugden 2001). ERKs and the upstream activators of
the MEKs were first identified from yeast (for review, see Levin & Errede 1995,
Herskowitz 1995; Fig. 4). The kinases in all levels are activated by Gq/G11 protein
29
coupled receptors and PMA in the heart (Bogoyevitch et al. 1994). Activation of c-Raf
and A-Raf occurs in response to PMA, ET-1 and PE, involving activation of PKC
(Bogoyevitch et al. 1995). In fact, several studies have found ERK as a downstream
target of PKC (Goldberg et al. 1997, Ho et al. 1998, Sugden & Clerk 1998b, Molkentin
& Dorn II 2001). PMA and ET-1 activate also MEKs and ERKs, whereas activation by αadrenergic agonist PE is less effective (Bogoyevitch et al. 1993, Bogoyevitch et al. 1994,
Bogoyevitch et al. 1996a).
Fig. 4. Schematic overview of MAP kinase cascades activated by diverse hypertrophic stimuli
and regulating phosphorylation and activation of numerous cardiac transcription factors
(modified from Sugden 1999). TNF-α, tumor necrosis factor-α; IL-1β, interleukin-1β; ATF-2,
activating transcription factor-2.
As noted, ERKs are activated by various neurohumoral stimuli, and also by myocyte
stretching in cultured cardiac myocytes and in vivo (Yamazaki et al. 1993, Sadoshima &
Izumo 1993, Yamazaki et al. 1995a). Transfection of constitutively active MEK1 into
cultured cardiac myocytes results in increased ANP promoter activity, whereas inhibition
of ERK cascade with dominant negative mutant of MEK1 abolishes PE induced ANP
promoter activity (Gillespie-Brown et al. 1995). Transgenic mice with cardiac restricted
expression of activated MEK1 demonstrated concentric hypertrophy with an
approximately 25% increase in heart to body ratio (Bueno et al. 2000, Bueno &
Molkentin 2002). Antisense oligonucleotide inhibition of ERK1 and ERK2 has been
shown to attenuate PE induced sarcomeric organization and increase in cell size, the
morphological changes seen in hypertrophy, as well as PE induced increase in ANP
promoter activity and ANP mRNA levels (Glennon et al. 1996). Since then, inhibitors of
ERK pathway, PD 98059 and U0126 have become commercially available (Favata et al.
1998, Dudley et al. 1995). ERK inhibition with PD 98059 has been shown to attenuate
the myofibrillar organization induced by PE or ET-1, supporting the role of ERKs in the
30
development of cardiac hypertrophy (Clerk et al. 1998b). However, contradicting data
also exist, showing that dominant-negative ERK1 or ERK2, or PD 98059 are not
sufficient to block PE induced ANP promoter activity or ET-1 induced increase in protein
synthesis (Post et al. 1996, Choukroun et al. 1998). Although the exact role of ERKs in
cardiac hypertrophy is not fully clear at present, there are convincing data indicating
involvement of ERK in mechanical stretch induced increase in c-fos and α-SkA gene
expression, and hypertrophic agonist induced increase in ANP promoter activity (Komuro
et al. 1991, Yamazaki et al. 1993, Yamazaki et al. 1995a, Gillespie-Brown et al. 1995,
Sadoshima & Izumo 1997). In fact, ERKs were the first MAPKs shown to phosphorylate
transcription factor c-Jun, although later studies proved JNKs to be principally
responsible for c-Jun phosphorylation (Pulverer et al. 1991, Karin 1995; Fig. 4). Recent
studies have also shown a role for ERKs in regulation of transcription factor GATA4
phosphorylation (Liang et al. 2001b; Table 2). Previously, Elk-1 transcription factor has
been shown to contain a phosphorylation motif for ERK2, and ERK pathway has also
been implicated in the regulation of Elk-1 dependent transcriptional activation (Yang et
al. 1998b, Babu et al. 2000).
Table 2. Nomenclature of MAP kinases and MAP kinase substrates (Kyriakis & Avruch
2001, Molkentin & Dorn II 2001, Wilkins & Molkentin 2002, Crabtree & Olson 2002).
SAPK, stress-activated protein kinase.
Name
ERK1
Alternate names
p44 –MAPK
ERK2
p42 –MAPK
JNK1
JNK2
JNK3
P38α
SAPK1c, SAPK-γ
SAPK1a, SAPK-α
SAPK1b, SAPK-β
SAPK2α, CSBP1
P38β
P38γ
P38δ
SAPK2β, p38-2
SAPK3, ERK6
SAPK4
Substrates
MAPKAP-K1, Elk-1, c-Jun, GATA4
MAPKAP-K1, Elk-1, c-Jun, GATA4
c-Jun, ATF-2, Elk-1, NFATc3
c-Jun, ATF-2, Elk-1, NFATc3
c-Jun, ATF-2, Elk-1, NFATc3
MAPKAP-K2/3, ATF-2, NFATc4,
MEF2
MAPKAP-K2/3, ATF-2, NFATc4
MAPKAP-K2/3, ATF-2
MAPKAP-K2/3, ATF-2
2.2.3.2 c-Jun N-terminal kinase
Molecular cloning of p54, JNK2, has revealed a family of SAPKs or JNKs, encoded by at
least three genes (Kyriakis et al. 1994, Derijard et al. 1994, Sluss et al. 1994; Table 2).
JNK activity is induced by a number of upstream kinases (Kyriakis & Avruch 2001; Fig.
4). The JNKs are serine/threonine kinases, which also, similarly to ERKs, require
tyrosine and serine/threonine phosphorylation for their activation (Kyriakis & Avruch
1990, Kyriakis et al. 1991). Interest in these kinases was heightened when they were
found to phosphorylate c-Jun, a component of the AP-1 transcription factor (Pulverer et
31
al. 1991, Karin 1995). JNKs are preferentially activated by cellular stresses, such as UVlight and osmotic stress, and by inflammatory cytokines, such as TNF-α and IL-1β
(Galcheva-Gargova et al. 1994, Hibi et al. 1993, Kyriakis et al. 1994, Derijard et al.
1994). In the heart, activation of JNKs, primarily JNK1 and JNK2, results from myocyte
stretch or activation of GPCRs (Ito et al. 1999, Cook et al. 1999, Choukroun et al. 1999).
A number of studies have implicated JNKs in the regulation of cardiac hypertrophy in
vitro and in vivo. Adenovirus mediated gene transfer of dominant negative mutant of
MKK4, an upstream kinase of JNKs, blocks ET-1 induced increase in protein synthesis,
sarcomere organization and ANP mRNA levels (Choukroun et al. 1998). Activated
MEKK1 has been shown to stimulate ANP reporter gene expression, while a dominant
negative MEKK1 mutant inhibits PE induced ANP expression (Ramirez et al. 1997,
Thorburn et al. 1997). Promoter studies have further revealed that activated MEKK1
induces β-MHC, ANP and α-SkA expression in cultured neonatal ventricular myocytes
(Bogoyevitch et al. 1996b). The results from studies using dominant negative or activated
MEKK1 are not clear, since MEKK1 has also been shown to induce activation of both
ERK and p38 MAPK pathways (Zechner et al. 1997, Thorburn et al. 1997). Crosstalk
between MAPK pathways is also known to exist at other levels, i.e. MEKK3, a
MAPKKK of ERK pathway, activating MAPKKs in ERK, JNK and p38 MAPK
pathways, as well as MKK4 activating JNKs and p38 MAPKs (Derijard et al. 1995,
Kyriakis & Avruch 2001; Fig. 4). Also, specific activation of p38α by adenovirusdelivered constitutively active MKK3β has been shown to result in potent inhibition of
the activity of ERK1,2 and its upstream activator MEK1,2 (Westermarck et al. 2001).
There are also contradictonary results concerning the role of JNKs in hypertrophic
signaling. Activated MEKK1 induced ANP expression was inhibited by activation of
JNK pathway, whereas in an earlier report MEKK1 induced ANP expression was JNK
dependent (Thorburn et al. 1997, Nemoto et al. 1998). Recently, studies with MEKK1
deficient mice showed that JNK signaling via MEKK1 is not required for aortic banding
induced cardiac hypertrophy and ANP expression, but protects the heart from apoptosis
(Sadoshima et al. 2002). Adenovirus mediated gene transfer of dominant negative MKK4
has been shown in vivo to inhibit pressure overload induced cardiac hypertrophy,
characterized by a decrease in LV thickness, LV to body weight ratio and ANP expression
(Choukroun et al. 1999). Primary targets for JNKs are transcription factors c-Jun, Elk-1
and ATF-2 (Gupta et al. 1995, Livingstone et al. 1995, Gupta et al. 1996, Sugden & Clerk
1998a; Fig. 4, Table 2). ATF-2 and c-Jun have been shown to form heterodimers that
transactivate at cAMP response element -like consensus sequences within various
promoters (van Dam et al. 1993, Sugden & Clerk 1998a). In the future, studies with
selective inhibitors are expected to further enlighten the JNK dependent mechanisms of
hypertrophic gene expression.
2.2.3.3 p38 mitogen-activated protein kinase
The p38 MAPK was also identified as a protein kinase cascade activated by IL-1β and
physiologic stress (Freshney et al. 1994). Like other MAPKs, p38 is activated by dual
32
tyrosine/threonine phosphorylation in catalytic domain (Freshney et al. 1994, Rouse et al.
1994). Four p38 isoforms have been described to date: p38α, p38β, p38γ and p38δ (for
review, see Kyriakis & Avruch 2001). The MAPKKs for p38 are MKK3 and MKK6,
which specifically activate p38 MAPKs (Derijard et al. 1995, Raingeaud et al. 1996). As
mentioned, MKK4, a MAPKK for JNK pathway, also phosphorylates and activates p38
in vitro (Derijard et al. 1995). The primary MAPKKK is MAPKKK5 (ASK1), which also
activates MKK4 and MKK7 in the JNK pathway (Kyriakis & Avruch 2001). MEKK1 is
also capable of inducing the p38 pathway, but not until MEKK1 expression robustly
exceeds that required for maximal JNK activation (Yan et al. 1994; Fig. 1). The p38
MAPKs are activated by a variety of stimuli, including chemical stress, physical stress,
osmotic stress and radiation, and by GPCR-agonist activation (Paul et al. 1997a).
In cardiac myocytes p38α and p38β are the most important isozymes, whereas p38γ
and p38δ are hardly detectable (Jiang et al. 1997). In the heart, p38 MAPKs are activated
by GPCR ligands, such as ET-1, PE, Ang II and by myocyte stretch (Sugden & Clerk
1998a, Liang et al. 2000). Activated p38 isoforms are shown to phosphorylate and
activate a number of transcription factors, including MEF2, ATF-2, ATF-6, nuclear factorkappaB (NF-κB) and Elk-1 (Raingeaud et al. 1996, Han et al. 1997, Thuerauf et al. 1998,
Liang & Gardner 1999, Yang et al. 1999a; Fig. 4, Table 2). In addition to transcription
factors, p38 MAPKs phosphorylate and activate several other protein kinases. These
include MAPK-activated protein kinases 2 and 3 (MAPKAP-K2 and MAPKAP-K3) and
p38-regulated/activated kinase, which phosphorylates the small heat shock protein
Hsp25/27 (Clifton et al. 1996; Table 2) (Clerk et al. 1998a, New et al. 1998). MAPKAP2 is expressed in adult cardiac ventricular myocytes as 66 kd alpha and 61 kd beta
subunits (World Health Organization 1998). Other known substrates for p38 MAPKs are
Mnk1 and Mnk2, and mitogen- and stress-activated protein kinases (Kyriakis & Avruch
2001).
In cultured cardiac myocytes p38 is rapidly activated by ET-1, PE and stretching of the
myocytes (Liang et al. 2000). In vivo pressure overload activates p38 MAPK, and
increased p38 activation has also been observed in human hearts with failure secondary
to coronary artery disease (Wang et al. 1998, Cook et al. 1999). In vitro overexpression
of activated MKK3 or MKK6 induces hypertrophic growth of myocytes and increased
expression of hypertrophic genes, such as ANP, BNP and α-SkA (Zechner et al. 1997,
Nemoto et al. 1998, Wang et al. 1998). Adenoviral overexpression of wild type p38β
results in increased cell size and ANP mRNA levels (Wang et al. 1998). Two relatively
specific inhibitors exist for p38α and p38β, SB203580 and SB202190 (Lee et al. 1993,
Goedert et al. 1997, Lee et al. 1994). In vitro pharmacological inhibition of p38 with SB
203580 (20 µM) blocks PE induced increase in ANP and BNP promoter activity, cell size
and sarcomeric organization (Zechner et al. 1997). SB 202190 at 20 µM concentration is
sufficient to suppress PE induced ANP promoter activity and ET-1 induced increase in
cell size and sarcomeric organization (Nemoto et al. 1998). Role of p38 is not fully clear,
since 10 µM concentration of SB 203580 failed to inhibit ET-1 and PE induced increase
in β-MHC expression and cell size, and only partially inhibited agonist induced
myofibrillar organization (Clerk et al. 1998b).
33
2.2.4 Phosphatases
2.2.4.1 Calcineurin
Calcium/calmodulin –dependent phosphoprotein phosphatase calcineurin, also known as
protein phosphatase-2B (PP2B), is a heterodimer composed of two subunits, calcineurin
A containing the catalytic site and calcineurin B containing the calcium binding
regulatory domain (Bueno et al. 2002a). There are three different mammalian genes
encoding calcineurin A, designated α, β and γ, of which calcineurin Aα and calcineurin
Aβ are present in the heart in multiple alternatively spliced forms (Bueno et al. 2002a).
Calcineurin B is encoded by two genes, each having alternatively spliced isoforms (Ueki
et al. 1992). Calcineurin physically interacts with NFATc family members and
dephosphorylates serine residues within these proteins (Rao et al. 1997). Once
dephosphorylated by calcineurin, NFATc transcription factors translocate to the nucleus
and directly activate immune response genes, usually in collaboration with AP-1 like
transcription factors (Flanagan et al. 1991, Loh et al. 1996a, Loh et al. 1996b). Nuclear
export of NFATs, in turn, is dependent upon rephosphorylation by glycogen synthase-3β
(GSK3β), JNK or p38 MAP kinases (Chow et al. 1997, Gomez et al. 2000, Neal &
Clipstone 2001). Using a yeast-two-hybrid screening for search of interacting factors for
transcription factor GATA4, a specific interaction was observed between zinc finger
domains of GATA4 with the DNA binding domain of NFATc4 (NFAT3). This interaction
was sufficient to induce synergic activation of BNP transcription (Molkentin et al. 1998).
Inhibition of calcineurin by cyclosporin A (CsA) or FK506 reduces PE and AngII induced
increase in cell size and sarcomeric organization (Molkentin et al. 1998). Transgenic
mice overexpressing the α-MHC driven catalytic subunit of calcineurin in the heart
demonstrate a vast increase in heart size and activation of expression of hypertrophic
genes β-MHC, α-SkA and BNP. Furthermore, treatment with CsA is sufficient to prevent
the dysfunction developed in the transgenic mice (Molkentin et al. 1998). At present
there are numerous studies supporting the hypothesis that calcineurin is involved in
hypertrophic signaling in cardiac myocytes (Wilkins & Molkentin 2002). Recent studies
have utilized naturally occurring inhibitors of calcineurin, so-called calcipressins, or
focused on inhibiting calcineurin involved transgenic overexpression of the calcineurin
inhibitory protein MCIP1 (myocyte-enriched calcineurin-interacting protein-1) to
genetically modify calcineurin mediated hypertrophic response (Taigen et al. 2000,
Görlach et al. 2000, Rothermel et al. 2001). Recent findings suggest proteolysis as a
mechanism regulating calcineurin phosphatase activity. Abundance of the calcineurin A
C-terminus containing the autoinhibitory domain was decreased in patients with
hypertrophic obstructive cardiomyopathy and aortic stenosis leading to increased NFAT2
electrophoretic mobility (Ritter et al. 2002, Williams 2002). In addition to NFATcs, also
MEF2 transcription factor is directly regulated by calcineurin (Passier et al. 2000).
Activation of calcineurin signaling is also associated with activation of other signaling
pathways, such as certain PKC isoforms, JNK and Akt (De Windt et al. 2000a, De Windt
et al. 2000b). Inhibition of calcineurin blocks β-adrenergic agonist induced activation of
ERKs and ET-1 gene expression (Zou et al. 2001, Morimoto et al. 2001).
34
2.2.4.2 Mitogen-activated protein kinase phosphatases
Other phosphatases suggested to participate in hypertrophic signaling in cardiac
myocytes involve mitogen-activated protein kinase phosphatases (MKPs). So far 11
MKPs have been identified, including MKP-1, MKP-2, hVH3/B23, MKP-3, MKP-X,
MKP-4, MKP-5, hVH-5/M3/6, PAC-1, MKP-6 and MKP-7 (Haneda et al. 1999, Masuda
et al. 2001). MKPs differ in terms of substrate specificity, tissue distribution, subcellular
localization and target gene regulation. In vitro, cultured neonatal cardiomyocytes
infected with a calcineurin-expressing adenovirus and stimulated with PE demonstrate
reduced p38 phosphorylation and increased MKP-1 protein levels. Inhibition of
calcineurin with CsA results in decreased expression of MKP-1 protein levels, further
supporting the interconnection between the pathways (Lim et al. 2001). Transgenic mice
overexpressing MKP-1 lack activation of all three MAPK branches, ERK, JNK and p38,
although p38 and JNK are preferred substrates over ERK (Franklin & Kraft 1997, Bueno
et al. 2001). These mice are characterized by reduced developmental hypertrophy
resulting in severe ventricular dilatation and neonatal lethality (Bueno et al. 2001). In
addition, Ang II treatment has been shown to increase MKP-1 mRNA levels, and thereby
negatively regulate MAPKs (Fischer et al. 1998, Hiroi et al. 2001).
Myocardial hypertrophy is associated with increased basal glucose metabolism,
characterized by increased expression of glucose transporter-1 mRNA. Transfection of
myocytes with MKP-3 has been shown to decrease TPA and PE induced increase in Glut1
mRNA levels (Montessuit & Thorburn 1999). MKP-3 shares 36% identity with MKP-1,
which efficiently blocks phosphorylation and activation of ERK2 (Muda et al. 1996).
MKP-2 can also be found in adult myocardium, where its levels are increased during
myocardial failure (Communal et al. 2002). Possible phosphatases worth further
investigation include PAC1 and M3/6, which have been shown to dephosphorylate p38 in
vivo (Mackay & Mochly-Rosen 2000).
2.3 Cardiac transcription factors
Cardiac gene expression is controlled via transcriptional and several posttranscriptional
mechanisms (Darnell, Jr. 1982). Several proteins inside the cell serve as regulatory
transcription factors, which recognize and bind specific DNA sequences in promoters
regulating gene transcription. Several transcription factors regulating the expression of
cardiac genes have been identified, many of those induced during cardiogenesis as well
as during hypertrophic cardiomyocyte growth (Srivastava 2001). To date, numerous
transcription factors have been identified within the promoters of cardiac genes that are
upregulated in response to increased hemodynamic load. Analysis of mechanisms
controlling the expression of natriuretic peptide genes, ANP and BNP, has revealed many
key regulatory elements, among which there are binding sites for transcription factors
GATA4, NFATc4 and Elk-1 (Nemer & Nemer 2001, Pikkarainen et al. 2003).
35
2.3.1 GATA family of transcription factors
Six different GATA factors have been identified, and they can be divided into two
subfamilies, GATA1, -2 and -3, and GATA4, -5 and -6. GATA1, -2 and -3 are
preferentially expressed in hematopoietic cells, whereas GATA4, -5 and -6 are expressed
in various tissues including the heart, liver, lung, gonad and gut (Arceci et al. 1993,
Kelley et al. 1993, Laverriere et al. 1994, Morrisey et al. 1996, Morrisey et al. 1997a,
Suzuki et al. 1996, for review, see Orkin 1998). Members of the GATA family of
transcription factors contain a highly conserved DNA binding domain with two zinc
fingers. This domain specifically interacts with DNA cis elements containing a consensus
(A/T)GATA(A/G) sequence. Mouse GATA4, -5 and -6 encode proteins of 48, 42 and 45
kDa, respectively (Arceci et al. 1993, Morrisey et al. 1996, Morrisey et al. 1997a). The
proteins are 85% identical to each other at the amino acid level within the DNA binding
region containing the zinc finger and basic regions (Molkentin 2000b). It has been
reported that only the C-terminal zinc finger and the adjacent basic domain are necessary
for specific DNA binding in vitro (Yang & Evans 1992, Omichinski et al. 1993, Visvader
et al. 1995). Using nuclear magnetic resonance-technique, the central and N-terminal
portions of GATA-1 were shown to interact with the major groove of DNA, whereas the
C-terminal zinc finger made site-specific interactions within the minor groove. In fact,
the N-terminal zinc finger of GATA1 has been shown to interact with adjacent GATA
DNA sequence elements and with other cooperating transcription factors (Trainor et al.
1996, Weiss et al. 1997). Deletion analysis of GATA4 revealed that the C-terminal zinc
finger is necessary and sufficient also for GATA4 DNA binding (Morrisey et al. 1997b).
Deletion analysis also suggested that GATA4 has a nuclear localization signal in the basic
domain adjacent to the C-terminal zinc finger (Morrisey et al. 1997b). On the other hand,
the N-terminus of GATA4 protein contains two separate transcriptional activation
domains, which are also partially conserved in GATA5 and GATA6, suggesting a similar
mechanism of transcriptional activation (Morrisey et al. 1997b).
Binding motifs for GATA-factors have been identified within the promoters of a
number of cardiac-expressed genes, including β-MHC (Hasegawa et al. 1997), α-MHC
(Molkentin et al. 1994, Huang & Liew 1997), cardiac troponin C (Ip et al. 1994), ANP
(Grepin et al. 1994), BNP (Thuerauf et al. 1994), cardiac troponin-I (Murphy et al. 1997,
Di Lisi et al. 1998, Bhavsar et al. 2000) and natrium-calcium exchanger (Nicholas &
Philipson 1999, Cheng et al. 1999). Studies with α- and β-myosin heavy chain promoters
have shown a preferential utilization of GATA4 over GATA6 (Charron et al. 1999). On
the other hand, the same study revealed that GATA6 participates in the regulation of
cardiac gene expression by regulating ANP and BNP promoter activities in cooperation
with GATA4 (Charron et al. 1999). Analysis of β-MHC promoter implicated that the
proximal region of the promoter containing the GATA binding sites directs gene
expression in response to pressure overload (Hasegawa et al. 1997). Similarly GATA4
has been shown to regulate pressure overload induced angiotensin type-1A receptor and
BNP promoter activity (Herzig et al. 1997, Marttila et al. 2001). More recently,
adenoviral overexpression of GATA4 in cultured neonatal cardiac myocytes enhanced
sarcomeric organization, induced an increase in cell surface area and resulted in a
significant increase in total protein accumulation (Liang et al. 2001a). Transgenic mice
36
with 2.5-fold overexpression of GATA4 within the adult heart demonstrate an increase in
heart to body weight ratio, histological features of cardiomyopathy, and activation of
hypertrophic genes, such as ANP, BNP and α-SkA, clarifying the functional role of
GATA4 in the development of hypertrophic phenotype (Liang et al. 2001a). In cultured
cardiac myocytes electrical pacing induced hypertrophy was associated with an increase
in GATA4 mRNA levels, further supporting the role of GATA4 in the development of
hypertrophic phenotype (Xia et al. 2000).
GATA4 participates in the regulation of cardiac gene expression in response to diverse
hypertrophic stimuli, including pressure overload, isoproterenol, PE, ET-1, Ang II and
PMA (Hasegawa et al. 1997, Morimoto et al. 2000, Hautala et al. 2001, Liang et al.
2001b, Kitta et al. 2001, Clement et al. 2002). Hypertrophic agonists activate GATAdependent gene expression by increasing GATA4 binding or transactivating activity, but
the signaling pathways involved are not clear (Liang & Molkentin 2002). Previous
studies in hematopoietic progenitor cells and in Cos cells have indicated an ERKdependent mechanism phosphorylating GATA2 (Towatari et al. 1995). In cardiac
myocytes PE induced ET-1 promoter activity was associated with phosphorylation of
GATA4 via ERK-dependent mechnism (Morimoto et al. 2000). The phosphorylation site
for ERK was later identified as Ser 105, which is situated at a site potentially regulating
GATA4 DNA binding. Phosphorylation of GATA4 by GSK3β results in cytosolic
localization and decreased activity of GATA4 (Morisco et al. 2001).
In addition to phosphorylation and increased gene expression, GATA4 is regulated by
interactions with other transcriptional cofactors. A number of transcription factors and
nuclear cofactors have been shown to interact with GATA4 to activate or repress cardiac
gene expression. GATA4 directly interacts with cardiac transcription factor NKX2.5 to
regulate expression of ANP and α-actin promoters (Durocher et al. 1997, Sepulveda et al.
1998, Lee et al. 1998). GATA factors also regulate developmental expression of NKX2.5,
suggesting a strengthened regulatory mechanism between these transcription factors
(Molkentin 2000b). GATA4 physically interacts with NFATc4 and MEF2 regulating
cardiac gene expression (Molkentin et al. 1998, Morin et al. 2000). Interaction of the Nterminal zinc finger of GATA4 with the transcriptional modifying protein FOG-2 has also
been described (Lu et al. 1999, Svensson et al. 1999, Tevosian et al. 1999).
2.3.2 NFATc family of transcription factors
A recently discovered intracellular pathway linking extracellular stimuli to hypertrophic
gene expression employs the calcineurin dependent downstream activator NFATc4
(Molkentin et al. 1998). Cardiac myocytes contain four members of NFATc proteins:
NFATc1 (also named as NFAT2 or NFATc), NFATc2 (NFAT1 or NFATp), NFATc3
(NFAT4 or NFATx) and NFATc4 (NFAT3). NFATs are cytoplasmic proteins, which are
translocated into the nucleus in response to stimulation that promotes Ca2+ mobilization
and activates calcineurin (Rao et al. 1997). Activated calcineurin interacts physically with
cytoplasmic NFAT proteins and dephosphorylates multiple serine residues within the Nterminal regulatory domains, resulting in nuclear translocation of the NFATc proteins
37
(Beals et al. 1997a, Aramburu et al. 1998, Aramburu et al. 1999). Phosphoserines within
these residues are believed to mask nuclear localization sequences (NLS).
Dephosphorylation exposes the NLS and results in rapid nuclear import of NFATc
proteins. In addition to calcineurin, constitutively activated Ras results in increased NFAT
activity and nuclear translocation of NFATc3 (Ichida & Finkel 2001). Ras appears to
function upstream of calcineurin, since CsA treatment blocks Ras induced NFATdependent gene expression and nuclear localization of NFATc4 (Ichida & Finkel 2001).
In the nucleus NFATc proteins can bind the DNA sequence GGAAAAT as monomers or
dimers, and they can also form complexes with other transcription factors to bind
composite DNA sequences in the regulatory regions of their target genes. In the nucleus
NFATc proteins can bind the DNA sequence GGAAAAT at the regulatory regions of their
target genes, usually in cooperation with AP-1-like factors binding to adjacent sites (Jain
et al. 1992, Boise et al. 1993, Macian et al. 2000).
Discovery of phosphoserine residues within N-terminus of NFATc controlling nuclear
export of NFATs gave rise to search for kinases controlling the translocation (Beals et al.
1997a). Purification of kinases able to phosphorylate these functional sites, but not other
sites within NFATc protein, led to identification of glycogen synthase-3 kinase (Beals et
al. 1997b). Later on it was discovered that GSK3β negatively regulates not only nuclear
translocation of NFATc, but also DNA binding activity of NFATc (Neal & Clipstone
2001). Other kinases shown to oppose NFATc localization into the nucleus include
protein kinase A, p38 MAPK and JNK, which control the export of NFATc2, NFATc3 and
NFATc4 (Beals et al. 1997b, Chow et al. 1997, Gomez et al. 2000, Yang et al. 2002). It
has also been found recently that activation of cGMP-dependent protein kinase type 1 by
NO/cGMP suppresses NFATc transcriptional activity, BNP induction and myocyte growth
in response to α1-adrenergic stimulation (Fiedler et al. 2002). In contrast to other known
NFATc kinases, Pim-1 kinase has been shown to enhance NFATc-dependent
transactivation by phosphorylating NFATc1 on several serine residues distinct from the
negative regulatory sites (Rainio et al., 2002). Evidence for both positive and negative
feedback loops regulating NFATC activity has also become evident. In T cells, NFATc1
expression is stimulated upon calcineurin-induced binding of NFATc2 to the NFAT site
within the NFATc1 promoter (Zhou et al. 2002). In muscle cells calcineurin and NFATc
activate MCIP1, an endogenous inhibitor of calcineurin, creating negative feedback
control (Yang et al. 2000). The upregulation of MCIP1 expression in cardiomyocytes is
mediated by a cluster of tandem NFAT binding sites in MCIP1 gene promoter (Yang et al.
2000). Recent studies indicate that MCIP1 is also a substrate for ERK and GSK3 (Vega et
al. 2002).
Involvement of NFATc signaling in the regulation of hypertrophic cardiomyocyte
growth was discovered by Molkentin et al., who found that transgenic mice with cardiacselective overexpression of either calcineurin or NFATc4 developed severe cardiac
hypertrophy (Molkentin et al. 1998). This suggests that, differently from the T-cells,
NFATc4 activation alone without other signaling elements can activate hypertrophy and
expression of its target genes. N-terminal deletion mutant of NFATc4 that lacks the sites
for calcineurin dephosphorylation is constitutively localized into the nucleus. The
observation that N-terminal deletion mutant of NFATc4 can induce cardiac hypertrophy
when overexpressed in vivo confirmed that dephosphorylation of NFATc4 is sufficient to
induce cardiac growth (Crabtree & Olson 2002). In addition to GATA4, MEF2 has also
38
been shown to associate with NFATc4 and control hypertrophic response genes (Blaeser
et al. 2000). NFATs together with cofactors have been implicated in the regulation of
BNP, β-MHC, ET-1 and MCIP1 gene expression (Wilkins & Molkentin 2002).
Calcineurin induced nuclear translocation of NFATc proteins is blocked by
immunosuppressive drugs FK506 or CsA, which block calcineurin activation.
Pharmacological studies employing the calcineurin inhibitors CsA or FK506 in different
rodent models of heart disease have produced largely controversial results, with some
studies reporting complete rescue from hypertrophy and others reporting either partial
effects or no effects at all (Olson & Molkentin 1999, Molkentin 2000a, Rothermel et al.
2001). It has also been reported that hypertrophy can be exacerbated by CsA treatment in
mice with a mutation in α-MHC gene resembling a mutation common in humans (Fatkin
et al. 2000).
Several genetic approaches involving calcineurin/NFATc signaling have been applied
in recent years. Transgenic mice expressing a constitutively active form of GSK3β under
control of α-MHC promoter suppressed calcineurin induced cardiomyocyte growth, with
reduced nuclear localization of NFATc (Antos et al. 2002). Overexpression of MCIP1 in
the heart under control of α-MHC promoter inhibits cardiac hypertrophy in response to
activated calcineurin, pressure overload and exercise (Rothermel et al. 2001). While
dephosphorylated NFATc4 is able to induce hypertrophy when overexpressed in the heart,
there is no evidence that hypertrophic gene expression requires direct binding of NFATc4
to target gene promoters.
2.3.3 The ETS-domain transcription factor family
The mammalian ETS-domain family of transcription factors is composed of nearly 30
members (Maroulakou & Bowe 2000). They were originally identified on the basis of a
region of primary sequence homology with the protein product of the v-ets oncogene
encoded by E26 (E twenty six) avian erythroblastosis virus (for review, seeKarim et al.
1990). The ETS-factors share a unique structure of the 85 amino acid ETS domain that
has been described as a winged helix-turn-helix (Donaldson et al. 1994). The DNA
binding of ETS-factors is also conserved, with all members recognizing a core enhancer
sequence (C/G)GA(A/T) (Woods et al. 1992, Gupta et al. 1998). In many cases ETSfactors regulate gene expression by co-operating with other transcription factors, such as
AP-1, and ternary complex factors (TCFs), such as SRF (Sharrocks et al. 1997).
There is a TCF subfamily of ETS-domain transcription factors, composed of three
members: Elk-1, SAP-1 and SAP-2 (Net) (Sharrocks et al. 1997). These proteins contain
four conserved domains; an N-terminal ETS DNA-binding domain; the B-box, which
binds directly to SFR; the D domain, which represents a docking site for MAPKs; and the
C-domain, which acts as an MAPK inducible transcriptional activator domain (Fig. 5)
(Treisman 1996, Sharrocks et al. 1997, Yang et al. 1998a, Li et al. 2000). In fact, the TCF
subfamily of ETS-factors represents a key nuclear target for MAPK pathways. There is
evidence that Elk-1 is activated by ERK, JNK and p38 MAPK pathways (Treisman 1996,
Yang et al. 2001). Elk-1 is phosphorylated at multiple sites within the C-terminal
39
transcriptional activator domain, resulting in both enhanced DNA-binding activity and
transactivation (Sharrocks et al. 1997). Phosphorylation of Elk-1 also results in change of
mobility of DNA-bound complexes, suggesting a conformational change in the complex.
Elk-1 has been proposed to exist in two conformations, such that either phosphorylation
by MAPKs or SRF binding to the B-box is believed to convert Elk-1 to an open
conformation that can efficiently bind DNA in both binary and ternary complexes (Yang
et al. 1999b). It has recently been shown that MAP kinase phosphorylation-dependent
activation of Elk-1 leads to activation of the co-activator p300, which may play a role in
the regulation of stress responsive immediate early genes (Li et al. 2003). There are also
studies demonstrating that interaction of Elk-1 with other transcription factors, such as
histone deacetylase-1 and Sp1, can lead to transcriptional repression (Li et al. 2000, Yang
et al. 2001).
SRF
A
SRF
CD
A
c-fos
Fig. 5. A model illustrating the possible role of Elk-1 and SRF in the regulation of c-fos gene
expression (modified from Drewett et al. 2000). A, ETS domain; C, carboxyl-terminal
regulatory domain, D, MAPK docking site of Elk-1.
Cardiac myocytes represent several targets for ETS factors. Pressure overload is known
to induce expression of c-fos, a member of the AP-1 transcription factor complex.
Deletion and point mutations in the SRE of the c-fos promoter result in loss of pressureinduced reporter gene expression, indicating that the SRE is necessary for pressure
response (Aoyagi & Izumo 1993). The minimal c-fos promoter consisting of SRE alone is
sufficient to confer pressure responsiveness, confirming that the pressure response
element coincides with the SRE (Aoyagi & Izumo 1993). p38 MAPK mediated Elk-1
binding on ETS binding sequence (EBS) within the BNP promoter has recently been
shown to regulate ET-1 induced BNP gene expression (Pikkarainen et al. 2003). The two
palindromic EBSs within the α-MHC promoter have been shown to act as strong
repressor elements for α-MHC gene expression (Gupta et al. 1998). In cultured chick
ventricular myocytes Ets-1 has been shown to participate in the regulation of inducible
nitric oxide synthase (iNOS) gene expression (Takahashi et al. 2001). Several cardiac
genes contain potential binding sites for ETS-factors in their promoters, including α-SkA,
cardiac α-actin, β-MHC, MLC-2, cardiac troponin T, cardiac troponin C and cardiac
troponin I, which therefore serve as potential targets for ETS-factors (Gupta et al. 1998).
40
2.3.4 HIF-1
Hypoxia-inducible factor (HIF)-1 is a heterodimeric DNA binding transcription factor
consisting of two subunits, 120 kDa HIF-1α and 91- to 94 kDa HIF-1β (Wang et al. 1995,
Bertges et al. 2002). HIF-1β can dimerize with several different basic helix-loop-helix –
PAS proteins, whereas HIF-1α is the oxygen regulated subunit responsible for HIF
activity (Wang et al. 1995, Semenza 1999). The regulation of HIF involves changes at
transcription level as well as posttranscriptional and posttranslational alterations in
response to hypoxia (Jung et al. 2002). The most characteristic regulatory mechanism for
HIF is its protein stabilization under hypoxic conditions, and its rapid degradation on
reoxygenation. Hydroxylation of a proline residue within HIF-1α degradation domain by
HIF1-α prolyl-hydroxylase under hypoxic conditions has been shown to regulate HIF1-α
stability and HIF transcriptional cascade (Jaakkola et al. 2001).
Under normoxic conditions HIF-1 activity is mainly regulated by ubiquitination by
multiple domains within the HIF-1 protein. In addition, there is an increasing body of
evidence indicating that also changes in phosphorylation of HIF-1 affect HIF-1
expression and transactivation of target genes. HIF-1 serves as a substrate for various
kinase pathways including PI3K, ERK, JNK and p38 MAP kinase (Minet et al. 2001). It
has been reported that ectopically overexpressed c-Jun cooperates with HIF-1 to regulate
hypoxic response element dependent reporter gene expression (Alfranca et al. 2002).
HIF-1 preferentially binds to a TACGTGCT motif that first identified in 3’ enhancer
element of erythropoietin gene (Semenza & Wang 1992). Functional HIF-1 binding sites
have since been identified within promoters of a number of hypoxia responsive genes,
such as vascular endothelial growth factor, glucose transporter-1 and lactate
dehydrogenase A (Bertges et al. 2002). In cardiac myocytes HIF-1 has been shown to
regulate the expression of AM, ET-1, iNOS and vascular endothelial growth factor
(Cormier-Regard et al. 1998, Jung et al. 2000, Kim et al. 2002, Kakinuma et al. 2002). A
recent study also revealed binding sites for HIF-1 in rat ANP promoter and showed that
overexpression of HIF-1alpha was sufficient to enhance ANP promoter activity (Chun et
al. 2003).
3 Aims of the research
The aim of the present study was to evaluate the role of GATA4 transcription factor in
hypertrophic cardiomyocyte growth and gene expression, and to study the mechanisms
regulating GATA4 function in response to GPCR stimulation. The second purpose was to
study the roles of MAPK and PKC signaling pathways in regulating GPCR agonist and
stretch induced cardiomyocyte growth and gene expression, and to identify nuclear factor
elements targeting the regulatory modules of load induced cardiac genes.
Specifically the aims were:
1. To study the specific role of GATA4 binding on BNP gene expression and
hypertrophic cardiomyocyte growth.
2. To define the role of MAPK pathway kinases in the regulation of ET-1 induced
GATA4 binding to BNP gene promoter and GATA-dependent BNP gene expression.
3. To study the role of PKCα on PE and ET-1 induced hypertrophic cardiomyocyte
growth and gene expression.
4. To characterize the role of ERK and p38 MAPK in regulating hypertrophic gene
expression in response to increased load in isolated rat atrium.
4 Materials and methods
4.1 Materials
The chemicals and supplies used in this study were: formaldehyde and guanidine
isothiocyanate (Fluka Chemie AG, Buchs, Switzerland), CsCl (Serva Feinchemica Gmbh
& Co, Heidelberg, Germany), LiCl (JT Baker Chemicals BV, Holland), agarose NA
(Pharmacia LKB Biotechnology, Uppsala, Sweden), Hybond N+ nylon membrane
(Amersham Life Science, Buckinghamshire, UK), Quick Prime Kit (Pharmacia, Sweden),
ET-1 (Phoenix Pharmaceuticals Inc., CA, USA and Peninsula Laboratories, Belmont, CA,
USA) and bovine myelin basic protein (MBP) (Upstate Biotechnology, Lake Placid, NY).
[γ-32P] ATP, [α-32P] dCTP, [3H] leucine, ECL plus Western blotting reagent, Hyperfilm
MP and p42/44 (ERK) kinase enzyme assay system were purchased from Amersham
International (Amersham, Bucks, UK). p38 In vivo Kinase Assay Kit (MercuryTM) was
from Clontech (Palo Alto, CA). Luciferase and β-galactosidase reagents were purchased
from Promega (Madison, WI), Fugene 6 transfection reagent from Roche Molecular
Biochemicals (Mannheim, Germany), DOTAP from Boehringer Mannheim (Mannheim,
Germany) and TFX-50 from Promega (Madison, WI). PhosphoPlus p38 MAP Kinase
Antibody Kit was purchased from New England Biolabs Ltd (Hitchin, Hertfordshire,
UK). GATA4, GATA5, GATA6 and HIF-1 polyclonal antibodies were from Santa Cruz
Biotechnology (Santa Cruz, Ca). Anti-phosphoserine antibody and anti-phosphotyrosine
antibody were from Zymed laboratories Inc (San Francisco, Ca). Anti-phosphothreonine
antibodies were obtained from Alexis Corporation (San Diego, Ca), Zymed laboratories
Inc (San Francisco, Ca) and Santa Cruz Biotechnology (Santa Cruz, Ca). pCMV-Flagp38α, pCDNA-Flag-JNK1 and pCDNA3-Flag-JNK1(APF) plasmids were kind gifts
from Dr. R.J. Davis (Howard Hughes Medical Institute, University of Massachusetts
Medical School). pMT2-mGATA4 plasmid was a gift from D.B. Wilson (Dept of
Pediatrics, St. Louis Children’s Hospital). pCMV-MEK1 and pCMV-MEKK1 plasmids
were purchased from Clontech (Palo Alto, CA). pUC19 plasmid was obtained from New
England Biolabs Ltd (Hitchin, Herts, UK). Other chemicals were from Sigma.
43
4.2 Experimental animals
Male 2-to-4-day-old and 8-week-old Sprague-Dawley (SD) rats were from the Center for
Experimental Animals at the University of Oulu, Finland. The animals were housed in
plastic cages with free access to tap water and normal rat chow in a room with controlled
40% humidity and temperature of 22 ºC, where a 0600 h on, 1800 off environmental light
cycle was maintained. The maintenance diet of the animals contained 0.65% sodium
chloride (NaCl). The Animal Use and Care Committee of the University of Oulu
approved the experimental design. The investigation conforms with the Guide for the
Care and Use of Laboratory Animals published by the US National Institutes of Health.
4.3 Experimental protocols
Study
I
II
Experimental model
Duration
Decoy oligonucleotide depletion of 15 min–48 h
GATA4 binding
Neonatal rat ventricular myocytes
PE and ET-1 induced hypertrophic 5 min–48 h
myocyte growth
Neonatal rat ventricular myocytes
III
Antisense oligonucleotide
inhibition of PKCα
Neonatal rat ventricular myocytes 5 min–48 h
IV
Left atrial wall stretch
Isolated perfused rat atrial
preparate
Neonatal rat atrial myocytes
35 min–24 h
Methods
Northern blotting
RIA
Gel mobility shift assay
Immunocytochemistry
Reporter gene assays
Western blotting
Immuno complex kinase assay
Gel mobility shift assay
Reporter gene assays
Northern blotting
RIA
RT-PCR analysis
Western blotting
Immunocytochemistry
Measurement of intracellular
calcium oscillations
Northern blotting
RT-PCR analysis
RIA
Western blotting
Kinase activity assays
Immunocytochemistry
Gel mobility shift assay
44
4.3.1 Neonatal rat cardiac myocyte cell culture (I–IV)
Ventricular and atrial cells were prepared from 2-to-4-day-old SD rats using the
collagenase dissociation method (Tokola et al. 1994). The hearts were perfused 6–8 times
for 5 minutes with disagregation medium (collagenase 1 g/l and CaCl2 25 µM in
phosphate buffered saline (PBS) at 37°C. The cells were filtered and washed twice in
DMEM/F-12 supplemented with 10% fetal calf serum and antibiotics (penicillinstreptomycin at 100 IU/ml). The cells were preplated onto 100 mm culture dishes for 15–
45 min at 37°C in humidified air with 5% CO2. The nonattached cells were collected and
plated at the density of 2 × 105 /cm2 onto Falcon wells from 15 to 60 mm in diameter.
Following 16-hour incubation, the myocytes were subjected to liposome-mediated
transfection with FuGENE 6, DOTAP (Boehringer Mannheim, Mannheim, Germany) or
TFX-50 (Promega, Madison, WI) for 6–8 hours. In reporter gene analysis experiments,
the transfection efficiency was controlled by co-transfecting the cells with Rous sarcoma
virus or cytomegalovirus (CMV) promoter driven β-galactosidase gene plasmids. The
antisense and decoy oligonucleotides (30 and 20 bases, respectively) used were
phosphorothioated to increase nuclease resistance and support RNase H cleavage of
hybridizing RNA. Sequence for GATA decoy oligonucleotide (ODN) was 5’TGTGTCTGATAAATCAGAGATAACCCCACC-3’ and for control ODN 5’TAAATTGGCCAAGTGTAGCTCCGTTTGTGA-3’.
Both
ODNs
with
their
complementary sequence ODNs were heated to +80°C and annealed at room temperature
for
2 h.
Sequence
for
antisense
PKCα
ODN
(As1)
was
5’ATTATCTCTGGTGATTTGGA 3’ targeting to 3’untranslated sequence on the rat PKCα
cDNA, and for scrambled ODN: 5’ GTGATATGTGCAGTTATTTC 3’. Two other
antisense PKCα ODNs used were 5’ TAAACGTCAGCCATGGTCCC 3’ targeting to
5’untranslated sequence (As2) and 5’ TTAGCGATGACCAGCTGATC 3’ targeting to
coding sequence (As3) on the rat PKCα cDNA. After transfection, cells were washed
twice with DMEM/F-12 and cultured in complete serum-free medium (CSFM), which
was DMEM/F-12 medium supplemented with 2.5 mg/ml bovine serum albumin, 1.0 µM
insulin, 5.64 µg/ml transferrin, 32 nM selenium, 2.8 mM sodium pyruvate, 1 nM T3 and
100 IU/ml penicillin-streptomycin. When appropriate, PE (100 µM) or ET-1 (100 nM)
were added to culture medium on the third day of culture. The protein kinase inhibitors
used (SB 203580, SB 202190 and PD 98059) were added to the culture medium 2–4
hours prior to exposure to PE or ET-1. Samples for RIA were taken before and after the
transfection, and at 8–24 hour intervals after the transfection. At the end of the
experiments, the cells were washed twice with ice cold PBS and either frozen at -70°C or
collected by scraping for further analysis.
4.3.2 Isolated perfused atrial preparations (IV)
Male SD rats weighing 290–400 g were used. The rats were decapitated, and the heart
from each rat was rapidly removed and placed in oxygenated (~10°C) buffer solution (in
mM: 113.8 NaCl, 28.6 NaHCO3, 2.5 CaCl2, 5.0 HEPES, 1.1 MgCl2, 1.2 KH2PO4, 4.7 L-
45
glutamic acid, 5.0 taurine, 5.0 creatine, 5.0 succinic acid, 5.0 glucose and insulin (0.06
nM); pH 7.4), which was also used at 37°C for superfusion of the atrium. The
experimental model used in this study was the isolated rat atrial appendix (Laine et al.
1994). The left atrial auricle was attached to one of the four ends of a cross-branch
polyethylene adapter, and the tissue was placed in a constant temperature (37°C) organ
bath. A tube was attached to the opposite end of the cross-branch adapter, and another
tube with a smaller diameter was inserted inside to carry the perfusate inflow into the
lumen of the atrium. For the outflow, another tube was attached to one of the cross
branches of the adapter. The same outflow tube was used to control the intra-atrial
pressure by adjusting the height of the end of the tube. The fourth cross branch of the
cross-branch adapter was connected to a pressure transducer (TCB 100, Millar
Instruments) to record the pressure in the lumen of the atrium. Inflow and outflow (2
ml/min) to both the atrial lumen and the organ bath at a constant temperature were
controlled by a peristaltic pump (7553-85, Cole-Parmer Instrument). Samples for RIA
were taken from the perfustate beginning from 20 minutes of the start of the perfusion.
For atrial stretch, the intra-atrial pressure inside the lumen was elevated from 2 to 8
cmH2O at 32 minute time point.
4.4 Isolation and analysis of cytoplasmic RNA (I, III–IV)
At the end of each experiment, the cultured cardiac myocytes were washed twice with ice
cold PBS and stored at -70°C. The atrial tissue was blotted dry, weighed, immersed in
liquid nitrogen and stored at -70°C until assayed. RNA was isolated by the guanidine
thiocyanate-CsCl method (Chirgwin et al. 1979). Northern blot hybridization, in which
the size and amount of specific mRNA molecules in total RNA preparations are
determined, was performed after isolation of RNA. For the RNA Northern blot analyses,
5–10 µg of samples of the RNA were transferred to Amersham Hybond N+ nylon
membranes. A 390-bp fragment of rat BNP complementary deoxyribonucleic acid
(cDNA) probe (Ogawa et al. 1991) (a generous gift from Dr. K. Nakao, Kyoto University
School of Medicine, Kyoto, Japan), full-length rat ANP cDNA probe (Flynn et al. 1985)
(a generous gift from Dr. P. L. Davies, Queen's University, Kingston, Ontario, Canada),
PCR amplified rat AM cDNA probe (nucleotides 287-736) (Romppanen et al. 1997), a
482 bp fragment of cDNA probe complementary to rat 18S ribosomal RNA (Lee et al.
1988b) were labeled with [32P]dCTP using T7 Quick Prime Kit (Pharmacia). cDNA
probes for rat α-SkA, rat β-MHC and rat c-fos were made with the RT-PCR technique.
Sequencing showed that the probes corresponded to bases 2950 to 3184 (GeneEMBL
access number v01218), 5794 to 5923 (x15939) and 231-1280 (x06769), respectively.
The membranes were hybridized overnight at +42ºC in 5 x SSC (saline sodium citrate, 1
x SSC = 0.15 M NaCl, 0.015 M trisodium citrate, pH 7), 0.5% sodium dodecyl sulfate
(SDS), 5 x Denhardt´s solution, 50% formamide and 100 µg/mL sheared herring sperm
DNA (Tokola et al. 1994). After hybridization, the membranes were washed in 0.1 x
SSC, 0.1% SDS three times for 20 min at +55ºC and exposed to Phosphor screen
(Molecular Dynamics, Sunnyvale, CA, USA) at room temperature. Phosphor screens
46
were scanned with Phosphor Imager (Molecular Dynamics) or Molecular Imager FX Pro
MultiImager System (Bio-Rad laboratories). The hybridization signals of AM, ANP, BNP,
c-fos, α-SkA and β-MHC mRNA were normalized to that of 18S RNA for each sample to
correct for potential differences in loading and/or transfer.
PKCα, β-MHC and HIF-1 mRNA levels were measured by quantitative reverse
transcription-PCR analysis with an ABI 3700 Genetic Analyzer using TaqMan chemistry
(Applied Biosystems, Foster City, CA) as previously described (Majalahti-Palviainen et
al. 2000). The RNA was extracted as previously described and a cDNA reaction was
performed according to the manufacturer’s protocol (Gibco BRL), after which mRNA
levels were measured by quantitative RT-PCR analysis. Forward and reverse primers for
PKCα
mRNA
detection
were
AGACCACAAATTCATCGCCC
and
CAAACCCCCAGATGAAGTCG, respectively. PKCα amplicon was detected using
fluorogenic probe 5'-FAM-CCCACCTTCTGCAGCCACTGCA-TAMRA-3'. The probe,
forward
and
reverse
primers
for
rat
β-MHC
were
5'-FAMTGTGAAGCCCTGAGACCTGGAGCC-TAMRA-3', GCTACCCAACCCTAAGGATGC
and TCTGCCTAAGGTGCTGTTTCAA, respectively. For rat HIF-1α, the probe, forward
and reverse primers were 5'-FAM-CCGGCGGCGAGAACGAGAAG-TAMRA-3',
GATTCGCCATGGAGGGC, and GACGTTCGGAACTCATCCTATTTT, respectively.
The results were normalized to 18S RNA quantified from the same samples using the
forward
and
reverse
primers
TGGTTGCAAAGCTGAAACTTAAAG
and
AGTCAAATTAAGCCGCAGGC, respectively. The probe for the 18S amplicon was 5'VIC-CCTGGTGGTGCCCTTCCGTCA-TAMRA-3'.
4.5 Radioimmunoassays (I, III–IV)
The medium samples were collected at 8–24 hour intervals in cell culture experiments
and the perfusate samples at 4 min intervals in isolated atrial perfusion experiments. The
immunoreactivity of ANP and BNP from cell culture medium and the immunoreactivity
of ANP from perfusate samples were determined directly from the sample. For isolated
atrial perfusate BNP radioimmunoassay, the 5 ml perfusate sample was extracted by SepPak C18 cartridges, lyophilized and redissolved to 500 µl of RIA buffer. The extracts and
unextracted perfusate and culture medium samples in duplicates of 100 µl were incubated
with specific rabbit BNP (Ogawa et al. 1991) and ANP (Jougasaki et al. 1995,
Vuolteenaho et al. 1985) antiserum. Synthetic rat BNP51-95 (BNP-45) and synthetic rat
ANP99-126, were incubated as standards. The tracers were prepared by chloramine-T
iodination of synthetic rat [Tyr0]-BNP51-95 and rat ANP99-126 followed by reverse phase
high performance liquid chromatography purification. After incubation for 48 hours at
+4ºC, the immunocomplexes were precipitated with sheep antiserum directed against
rabbit gammaglobulin in the presence of 500 µl of 8% Polyethylene Glycol 6000, pH 7,
followed by centrifugation at 3000 g for 30 min. The sensitivities of the BNP and ANP
assays were 2 fmol/tube and 1 fmol/tube, respectively. The intra- and inter-assay
variations were less than 10% and 15%, respectively. Serial dilutions of perfusate and
tissue extracts showed parallelism with the standards. The BNP antiserum did not
47
recognize ANP or CNP. The ANP antiserum recognized ANP and proANP with equal
avidity, but did not cross-react with BNP or CNP.
4.6 Reporter gene assays (I–II, IV)
Rat BNP promoter fragment was generated by PCR, with rat genomic EMBL3-λ-clone as
a template and using the following primers: sense 5´-GGGATTTGAACTCAGG -3´ with
KpnI, MluI-linker; antisense 5´-CACTAGCCTCTCAGCAACG -3´ with BamH1-linker.
Subsequently, PCR-product was digested with KpnI and BamH1 and cloned to KpnIBglII-site of pGL3-Basic vector (Promega) resulting in a (∆–5kbp/+4) BNP promoter
construct.
(∆–5kbp/+4) BNP-pGL3 construct was used to produce a (∆–538/+4) BNP-pGL3 by
nested deletion (Pharmacia Biotech). Site-directed mutations to two adjacent (–91 and
−80) GATA-sites of (∆–538/+4) BNP-pGL3 were prepared (Stratagene) and the resulting
construct is referred to here as BNP Gmut. The primers for mutagenesis were sense 5´GGCAGGAATGTGTCTTGAAAATCAGATGAAACCCCACCCCTAC-3´; antisense 5´GTAGGGGTGGGGTTGCATCTGATTTGCAAGACACATTCCTGCC-3´. A reporter
plasmid containing three GATA element consensus sequences linked to pXP-1 vector
(Grepin et al. 1994) was a kind gift from Dr. M. Nemer (Institut de Recherches Cliniques
de Montréal, Montreal, Canada).
4.7 Gel mobility shift assys (I–II, IV)
Nuclear extracts from cardiomyocytes were prepared as described previously (Schreiber
et al. 1989). Protein concentration from each sample was determined by using Bradford
assay (Bradford 1976) (Bio-Rad Laboratories). Double stranded-oligonucleotide
corresponding to GATA binding region (∆–68/–97) of rat BNP promoter was used for
analysis of GATA DNA binding activity and a previously described ODN for
measurement of Octamer-1 (Oct-1) DNA binding activity (Kemler et al. 1989; Table 3).
Both probes were sticky-end-labeled with [α−32P]dCTP by Klenow enzyme. For each
reaction mixture (20 µl) 6 µg of nuclear protein and 2 µg of poly(dI-dC) was used in a
buffer containing 10 mM HEPES (pH 7.9), 1 mM MgCl2, 50 mM KCl, 1mM
dithiothreitol (DTT), 1mM EDTA, 10% Glycerol, 0.025% NP-40, 0.25mM
phenylmethylsulfonyl fluoride (PMSF) and 1 µg/mL of each leupeptin, pepstatin and
aprotinin. Reaction mixtures were incubated with a labeled probe for 20 min followed by
non-denaturating gel-electrophoresis on 5% polyacrylamide gel. Subsequently, the gels
were dried and exposed in a PhosphorImager screen and analyzed with ImageQuant
(Molecular Dynamics) or Molecular Imager FX Pro MultiImager System (Bio-Rad
laboratories). To confirm DNA sequence specificity of the protein-DNA complex
formation, competition experiments with 10-, 50- and 100-molar excesses of nonradiolabeled ODNs with intact or mutated binding sites were performed. For competition
48
and supershift experiments, appropriate ODNs or antibodies (Santa Cruz Biotechnology)
were added to reaction mixture 20 minutes before addition of labeled probe.
Table 3. Probes for gel mobility shift analysis.
Probe
GATA (rat BNP)
NFAT (human BNP)
HIF-1 (mouse AM)
Oct-1 (human HH2A)
Sequence
for: TGTGTCTGATAAATCAGAGATAAC
rev: GGTGGGGTTATCTCTGATTTATCAGACACA
for: AGAGCTATCCTTTTGTTTTCCATCCTG
rev: CGGGCAGGATGGAAAACAAAAGGATAG
for: AGAGCCCGTGGCAAACGTGTTC
rev: CGGGGAACACGTTTGCCACGGG
for: GATCCGAGCTTCACCTTATTTGCATAAGCGATTGA
rev: GATCTCAATCGCTTATGCAAATAAGGTGAAGCTCG
4.8 Western blot analysis (II–IV)
Cells were washed twice with ice cold PBS and collected by scraping into 150–500 µl of
lysis buffer, which consisted of 20 mM Tris (pH 7.5), 150 mM NaCl, 1 mM EDTA, 1
mM EGTA, 1% Triton X-100, 2.5 mM sodium pyrophosphate, 1 mM βglycerophosphate, 1 mM Na3VO4, 1 µg/ml leupeptin, 1 µg/ml pepstatin, 1 µg/ml
aprotinin, 2 mM benzamidine, 1 mM PMSF, 2 mM and 50 mM NaF. The extracts were
further lysed with sonication and supernatant was collected after centrifugation. For
western blot analysis, cell lysates were matched for protein concentration (10–20 µg),
loaded on SDS-PAGE and transferred to nitrocellulose filters. The membranes were
blocked in 1–5% nonfat milk and then incubated with phospho-p38α, p38α, NFATc3,
NFATc4, phospho-Elk-1, Elk-1, GATA4, anti-serine, anti-threonine, anti-tyrosine, PKCα,
PKCβII, PKCδ, PKCγ or PKCε antibody overnight at 4°C. The filters were washed the
following day and incubated for 1 hour with an HRP conjugated anti-mouse, anti-rabbit
or anti-goat secondary antibody. The amount of protein was detected by enhanced
chemiluminescence.
4.9 Kinase activity assays (II, III)
Following treatment with appropriate agents, myocytes (approximately 1 × 106) were
washed with ice cold PBS at room temperature. Samples for PKC activity and ERK
activity assays were collected by scraping into 100 µl of lysis buffer containing 10mM
Tris (pH 7.5), 150 mM NaCl, 2 mM EGTA, 2 mM DTT, 1 mM Na3VO4, 10 µg/ml
leupeptin, 10 µg/ml aprotinin, 2 µg/ml pepstatin and 5 mM benzamidine. The extracts
were sonicated and supernatant was collected after centrifugation. 15 µl of protein extract
was incubated at 30°C for 15 minutes with 10 µl of substrate buffer containing specific
49
substrate peptide in the presence of 1 µCi (γ-32P) ATP. Each reaction was terminated and
blotted on separate peptide binding paper discs, which were washed repeatedly with 75
mM orthophosphoric acid. The incorporated radioactivity was measured with a
scintillation counter (Rackbeta II, LKB Wallac).
For immuno complex kinase assay endogenous p38 was immunoprecipitated with
specific antibody (New England Biolabs) at 4°C overnight, followed by protein GSepharose precipitation. The immunoprecipitates were washed three times with buffer
containing 50 mM Tris (pH 7.4), 150 mM NaCl, 5 mM EDTA, 25 mM βglycerophosphate, 25 mM NaF and 1% Triton X-100. Lysates were washed once more
with kinase buffer containing 25 mM Tris (pH 7.5), 5 mM β-glycerophosphate, 2 mM
DTT, 1.0 mM Na3VO4 and 10 mM MgCl2. The activity of the immuno complex was
assayed at 30°C for 15 minutes in 30 µl of kinase buffer in the presence of 2 µCi [γ-32P]
ATP and 20 µg of MBP as substrate (Kumar et al. 1997). The reactions were terminated
and the reaction contents electrophoresed on 15% SDS-polyacrylamide gels followed by
PhosphorImager or Molecular Imager FX Pro analysis to determine the phosphorylation
level of MBP. The effect of p38 inhibitor SB 203580 on p38 activity was measured by in
vivo kinase assay.
4.10 Calcineurin activity assay (III)
A specific kit was used for measurement of calcineurin activity (Biomol, Plymouth
Meeting, PA). According to the manufacturer’s instructions, the cells from neonatal rat
ventricular cell culture were washed twice with ice cold PBS and collected by scraping
into 300 µl of lysis buffer including a mixture of proteases inhibitors provided by
manufacturer. Cell lysis was further centrifugated for 45 min at 15000 g. The supernatant
was incubated with RII phophopeptide, a known substrate for calcineurin, and the free
phosphate released was detected based on the Malachite green assay (Martin et al. 1985).
In cellular extracts RII phosphopeptide is also cleaved by other phosphatases; i.e. PP1,
PP2A and PP2C. Hence, okadaic acid was used to inhibit PP1 and PP2A and, since EGTA
inhibits calcineurin, okadaic acid and EGTA were used to reveal the PP2C activity.
4.11 Cytochemistry (I, III)
For immunocytochemistry, cardiomyocytes were grown on glass coverslips. Cells were
washed in PBS, fixed in 4% paraformaldehyde containing 0.2% Triton X-100, washed
three times in PBS and once with pure ethanol in −20°C. Cells were next placed in ice
bath, washed three times with cold PBS and blocked in PBS containing 10% FBS and
0.02 M glycine. Sarcomeric organization was visualized by labeling α-actin filaments
with Alexa Fluor 568 Phalloidin (Molecular Probes, Eugene, OR). PKCα was assessed
using antibody against PKCα (Upstate Biotechnology, Lake Placid, NY) at a dilution of
1:400 in blocking solution. Subsequently, the cells were washed twice with cold PBS-
50
glycine, and anti-mouse FITC-conjugated antibody (Santa Cruz Biotechnology, Santa
Cruz, CA) was added at a dilution of 1:200 in blocking solution. Finally cells were
washed twice with PBS, once with dH2O and subjected to fluorescence microscopy.
For calcium concentration transient measurements and transfection efficiency
experiments the cardiac myocytes were cultured on glass bottom wells and, as desired,
loaded with Fluo-3 (10 µM, Molecular Probes) or transfected with complexes of 3´fluorescein isothiocyanate (FITC) labeled ODNs and FuGENE 6 as described above. The
intracellular calcium transients and localization of 3’FITC labeled ODNs in
spontaneously beating ventricular myocytes was examined using an LSM 510 laser
scanning confocal microscope (Zeiss, Thornwood, NY) equipped with argon laser and
attached to a Zeiss Axiovert 100 TV microscope (Williams 1990). Differential
interference contrast and green fluorescence channels were used.
4.12 Protein synthesis (I–III)
[3H]leucine-incorporation and total cellular protein content were measured from cells
cultured and transfected in 24-well plates. On the third day in culture, medium was
replaced with CSFM supplemented with [3H]leucine (5 µCi/mL) and when appropriate,
with ET-1 (100 nM) or PE (100 µM). After 24 hours cells were lysed and processed for
measurement of incorporated [3H]leucine (Amersham-Pharmacia) by liquid scintillation
counter (Rackbeta II, LKB Wallac) and total cellular protein content by Bradford assay
(Bradford 1976) (Bio-Rad Laboratories).
4.13 Statistical analysis
The results are expressed as mean±standard error of mean (SEM). Differences between
data groups were evaluated for significance using Student’s t test of unpaired data or oneway analysis of variance and Bonferroni’s post-test. Repeated measures ANOVA was
used for multivariate analysis. Differences at the 95% level were considered statistically
significant.
5 Results
5.1 Role of GATA4 in the regulation of BNP gene expression (I)
5.1.1 GATA4 DNA binding activity
The proximal (∆–68/–97) region of the rat BNP promoter contains two adjacent GATA
sites, which control cardiac-specific expression of BNP (Grepin et al. 1994). ET-1
increased GATA binding to (∆–68/–97) BNP promoter within 30 minutes, sustained for 3
hours. As a control for the specificity for GATA DNA binding activity, Oct-1 DNA
binding activity remained unchanged. Supershift analysis with specific antibody to
GATA4 confirmed the presence of GATA4 in the BNP promoter binding complex.
Specific antibodies of GATA5 and GATA6 had no effect on the electrophoretic mobility
of the BNP promoter binding protein complex.
In order to block GATA DNA binding, the proximal (∆–68/–97) rat BNP promoter
sequence was used to generate a 30 bp GATA4 factor binding cis-element decoy ODN.
Transfecting the GATA decoy ODN to cardiac myocytes abolished both basal and ET-1induced GATA4 DNA binding (> 95%) at 24 hours after transfection when compared to
cells transfected with control ODN, and similar downregulation of GATA binding was
detected at 48 hours after transfection. In vitro GATA4 DNA binding was totally
abolished by preincubation of nuclear extracts of cardiac myocytes with 50-molar excess
amount of the GATA decoy ODN but not with 50-molar excess of control ODN. Neither
of the ODNs competed with Oct-1 DNA binding. Blockade of GATA element binding
factors was specific and had no effect on Oct-1 DNA binding in ODN treated
cardiomyocytes. A positive control plasmid containing three GATA response elements
was used to verify the specific interaction of GATA with its target promoter. Cotransfection with GATA decoy ODN significantly decreased GATA-dependent promoter
activity, whereas control ODN was without effect. When GATA4 binding to rat ANP
promoter was studied, GATA4 also bound on (∆-109/-131) of rat ANP promoter, and the
52
DNA binding was totally abolished by preincubation of nuclear extracts with 50-molar
excess amount of the GATA decoy ODN, but not with control ODN.
5.1.2 Effect of GATA decoy treatment on ANP and BNP
ir-ANP
5
Relative to control
Relative to control
To study the effect of GATA decoy ODN treatment on the hypertrophic pattern of gene
expression, the expression of cardiac natriuretic peptides was evaluated. GATA decoy
ODN treatment significantly decreased basal secretion and gene expression of ANP and
BNP in cardiac myocytes (Fig. 6). Interestingly, ET-1 induced secretion of ANP and BNP
was independent of blocked GATA4 DNA binding, and similarly the increase of ANP and
BNP mRNA by ET-1 remained unaffected at 24 hours.
4
†
†
3
2
*
1
0
ET-1
control ODN
GATA decoy ODN
+
-
+
+
-
-
+
-
+
+
ir-BNP
10
ET-1
control ODN
GATA decoy ODN
†
†
8
6
4
2
0
*
+
-
+
+
-
-
+
-
+
+
Relative to control
BNP promoter activity
3
†
†
†
2
1
*
0
-5kb -538bp
-538bp
Gmut
Fig. 6. Effect of GATA decoy ODN treatment on basal end ET-1 induced secretion of ir-ANP,
ir-BNP and activation of a 5kb and 538bp rat BNP promoters in cultures neonatal ventricular
myocytes. Open bars, untreated cells; solid bars, ET-1 treated cells. Results are mean ± SEM.
*P < 0.01, †P < 0.001 compared with untreated control cells.
To study the rat BNP promoter region we introduced 5kb and 538bp BNP promoter
luciferase constructs to the myocytes. Deletion of a –5kb to –538 bp upstream region of
the BNP promoter did not affect basal promoter activity or ET-1-inducibility of the gene.
The mutation of two proximal GATA binding sites at –91bp and –80 bp of the 538bp
BNP promoter significantly decreased basal promoter activity, but it had no effect on ET1 induced promoter activity (Fig. 6).
53
5.1.3 Effect of GATA decoy on protein synthesis and sarcomere
assembly
Hypertrophic cardiomyocyte growth in response to stretch and exposure to various
agonists involves increased protein synthesis and increased organization of contractile
proteins to sarcomeric structures (Shubeita et al. 1990). To study the effect of GATA
decoy ODN treatment on ET-1-induced de novo protein synthesis, incorporation of 3Hlabeled leucine into the myocytes was studied. We found that GATA decoy ODN
treatment had no effect on basal or ET-1 induced protein synthesis and total cellular
protein content. Neither did GATA decoy ODN treatment affect ET-1 induced increased
organization of sarcomeres.
5.2 Role of MAP kinases in GATA4 activation and BNP gene
expression (II)
5.2.1 Activation of ERK and p38 MAP kinase by ET-1
p38 MAP kinase is activated in neonatal rat ventricular myocytes by various extracellular
stimuli, such ET-1 and PE (Clerk et al. 1998b, Liang et al. 2000). Western blot analysis
was used to confirm p38 activation by ET-1. ET-1 induced phosphorylation of p38 was
noted at 15 minutes. The kinetics of p38 activation was measured by immuno complex
kinase assay. Endogenous p38 was immunoprecipitated with anti-p38 antibody and its
activity was measured using MBP as a substrate. ET-1 induced a rapid increase in p38
activity, which was maximal at 15–20 minutes. In vivo kinase assay, which uses ATF-2 as
a substrate, was used to verify the inhibition of p38 by SB 203580 in cardiac myocytes.
Treatment with ET-1 (100 nM) for 24 hours led to a 3.4-fold increase of p38 activity, and
activity was totally inhibited by p38 inhibitor SB 203580 (20 µM), which also decreased
basal activity of p38 MAP kinase by 50%. In contrast, treatment of myocytes with a
potent MEK1 inhibitor PD 98059 increased basal p38 activity, but had no effect on ET-1
induced p38 activity.
Activation of ERK by GPRC agonists has been previously established (Sugden &
Clerk 1997). ERK activity was measured by an assay which measures transfer of a
phosphate group to a peptide highly selective for ERK. As reported previously
(Bogoyevitch et al. 1993, Liang et al. 2000), ET-1 at the concentration of 100 nM was a
strong activator of ERK. The response was maximal at 5 minutes and declined to almost
basal level within 35 minutes. MEK1 inhibitor PD 98059 (20 µM) was sufficient to
inhibit ET-1 induced ERK activation by 80% measured at 5 minutes.
54
5.2.2 Effect of ERK and p38 inhibition on ET-1 induced protein
synthesis
To examine whether blockade of p38 MAPK or ERK with specific inhibitors is sufficient
to attenuate ET-1 induced protein synthesis, incorporation of 3H-labeled leucine in
cardiac myocytes was examined. Treatment of myocytes with SB 203580 or PD 98059 at
the dose of 20 µM had no effect on basal protein synthesis. ET-1 induced 2.5-fold
increase in [3H]leucine incorporation was totally abolished by p38 MAP kinase inhibition
with SB 203580 (20 µM), whereas ERK inhibition with PD 98059 (20 µM) had no effect.
5.2.3 Regulation of GATA4 by MAP kinases
To examine GATA4 activation by ET-1 and the role of MAP kinases in the activation of
GATA4, gel mobility shift assay was applied. Activation of GATA binding to BNP
promoter occurred rapidly in response to ET-1 (100 nM). ET-1 stimulated GATA4
binding was detectable in 15 minutes and was maximal in 60 minutes. GATA4 binding
remained upregulated for 3 hours, and supershift analysis revealed that GATA4 was the
major cardiac nuclear factor that binds to the BNP GATA site. Mutation of either of the
GATA-binding sites showed that GATA4 binding was equal to both sites, but the binding
activity was reduced to about half of that observed with a probe having both sites intact.
Next the myocytes were pretreated with p38 and ERK inhibitors (SB 203580 and PD
98059, respectively), or the cells were transfected with dominant negative form of JNK
and then subjected to ET-1 treatment. The induction of GATA4 binding to BNP gene was
completely inhibited by p38 inhibitor SB 203580, and moreover, this inhibition of
GATA4 binding was dose-dependent (Fig. 7). Inhibition of ERK pathway with PD 98059
had no effect on ET-1 induced increase in GATA4 DNA binding, but basal GATA4
binding activity was significantly decreased (Fig. 7). Inhibition of JNK pathway with
dominant negative form of JNK had no effect on basal or ET-1 induced GATA4 DNA
binding.
To further elucidate the role of p38 MAP kinase in the induction of GATA4 DNA
binding, the p38 protein levels were increased by transfecting the myocytes with a CMV
promoter driven plasmid overexpressing p38α. ERK and JNK pathways were studied
similarly by using CMV promoter driven plasmids overexpressing MEK1 and MEKK1.
Myocytes transfected with pUC-19 were used as controls. Overexpression of p38
substantially evoked GATA4 binding to BNP gene as compared to control plasmid.
Treatment with p38 inhibitor SB 203580 completely abolished the effect. Inhibition of
ERK with PD 98059 (20 µM) resulted in a modest decrease in p38 induced GATA4
binding to BNP gene. Overexpression of MEK1 or MEKK1 had no effect on GATA4
DNA binding.
**
4
3
#
2
##
1
SB (uM)
ET-1 100 nM
0
5
20 0
5
20
- - - + + +
GATA-binding
(Relative to basal)
GATA-binding
(Relative to basal)
55
**
4
3
****
2
*
1
PD (uM)
ET-1 100 nM
0
5
20
0
5
20
- - - + + +
Fig. 7. Effect of p38 and ERK inhibitors (SB 203580 and PD 98059, respectively) on ET-1
induced GATA4 binding activity. The results are mean ± SEM. **P < 0.001, *P < 0.05
compared with untreated control cells, #P < 0.05, ##P < 0.001 compared with ET-1 treated
cells.
To examine whether p38 induced increase in GATA4 DNA binding activity was due to
changes in phosphorylation of GATA4, the myocytes were transfected with plasmids
overexpressing p38α, MEK1, MEKK1 or pUC19 (control). Subsequently, GATA4
phosphorylation was examined by Western blot analysis. Immunoblotting with GATA4
antibody showed that GATA4 protein levels were unaffected. Overexpression of MEK1
and p38α exhibited a marked increase in serine phosphorylation of GATA4, whereas
overexpression of JNK pathway (MEKK1) had no effect. p38 induced serine
phosphorylation of GATA4 was inhibited by p38 inhibitor SB 203580 and also by ERK
inhibitor PD 98059 consistently with the finding that p38α induced GATA4 binding was
also suppressed by PD 98059. Forced expression of p38, MEK1, or MEKK1 did not
induce threonine phosphorylation or tyrosine phosphorylation of GATA4.
To further investigate the role of MAP kinases in the regulation of GATA4, COS-1
cells transiently expressing GATA4 were cotransfected with plasmids overexpressing
p38α, MEK1 or JNK1. Control cells, cotransfected with pUC19, showed modest GATA4
binding activity to BNP promoter. p38α overexpression resulted in a 4-fold increase in
GATA4 binding activity, whereas MEK1 or JNK1 overexpression had no effect on
GATA4 binding to BNP gene promoter. Oct-1 binding activity was not affected by
transient expression of different plasmids.
5.2.4 p38 MAP kinase regulation of GATA-dependent promoter
To test whether p38 overexpression is sufficient to stimulate BNP promoter activity, the
myocytes were co-transfected with (∆–534bp/+4bp) BNP promoter plasmids and p38α
expression plasmid or pUC19 plasmid (control). p38α overexpression resulted in a 4-fold
increase in rat BNP promoter activity (Fig. 8). The mutation of two proximal GATA
binding sites at –91 bp and –80 bp of the rat BNP promoter totally abolished p38 induced
56
Fold activation
(Relative to basal)
increase in promoter activity. Cotransfection with a plasmid expressing either MEK1 or
MEKK1 induced both the BNP and the mutated constructs similarly.
6
*
5
4
3
2
#
1
p38
-
+
-534BNP
-
+
-534(Gmut)BNP
Fig. 8. p38 MAP kinase regulation of a luciferase linked 534bp rat BNP promoter with GATA
binding sites intact or mutated (Gmut). The results are mean ± SEM. *P < 0.05 compared
with basal -534BNP promoter activity, #P < 0.05 compared with p38 induced -534BNP
promoter activity.
5.3 PKCα in the regulation of cardiomyocyte hypertrophy (III)
5.3.1 Antisense inhibition of PKCα
Transfecting cardiac myocytes has been found difficult compared to many other cell
types. In the current study, three different cationic lipids were studied to optimize the
transfection method and to ensure that the lipid treatment did not interfere with the
results. In the experiments, highest transfection efficiency was achieved with FuGENE 6,
which also showed least toxicity analyzed with microscope imaging of the cells. To
examine the transfection efficiency of the antisense PKCα ODNs using the cationic
liposome delivery system, fluorescent-tagged ODNs (500 nM) corresponding to PKCα
were added with FuGENE 6, and the cells were subjected to confocal microscopy.
Increased accumulation of fluorescein-tagged particles inside the myocytes revealed that
both antisense and scrambled ODNs were effectively delivered into the cells with
FuGENE 6. The proportion of fluorescent-tagged ODN transfected myocytes in the
culture was ~60–80%. The strongest fluorescence was seen in the areas around the
nucleus. On the second day after the transfection, fluorescence staining was decreased,
but still substantial. To verify that the results were not influenced by the lipid treatment,
the experiments were repeated using also DOTAP and TFX-50 as lipids. The most
effective transfection ratio of ODN and FuGENE 6 was 1:2, whereas the most effective
ratios for both DOTAP and TFX-50 were 2:3 (ODN:lipid). The transfection efficiencies
with DOTAP and TFX-50 were considerably lower than with FuGENE6. Additionally,
57
light microscope imaging of the cells revealed striking morphological changes in DOTAP
treated cells. In the absence of cationic liposome carrier, only minimal transfection
efficiency was observed.
Several ODNs, each 20 bases in length, were designed to hybridize to different regions
of rat PKCα mRNA. Treatment of myocytes with the PKCα antisense ODN (0.5 µM)
targeted to 3’untranslated sequence (As1) on rat PKCα cDNA reduced PKCα mRNA
levels by 50% compared with scrambled ODN. The levels of 18S RNA were unaffected
by the PKCα antisense ODN treatment. In agreement with changes in PKCα mRNA
levels, treatment with antisense PKCα ODNs reduced PKCα protein levels by more than
60%. Dose response studies revealed that maximal reduction in PKCα protein levels was
achieved at 0.5 µM, whereas control ODN with the same base composition as the
antisense ODN but with a scrambled sequence had no effect. Western blot analysis with
antibodies selective to PKCδ, PKCε, PKCζ and PKCβII showed that antisense PKCα
treatment had no effect on other PKC isozymes. In untreated control cells PKCα staining
was diffuse around the cell, and treatment with PE (100 µM) for 5 minutes translocated
PKCα to the perinuclear region. In antisense PKCα ODN treated cells, fluorescence
staining for PKCα was dramatically decreased, whereas in control ODN treated cells a
significant amount of PKCα was seen around the nucleus.
PKC activity was studied by measuring the transfer of a phosphate group to a peptide
substrate highly selective for protein kinase C. PE treatment evoked a 2.8-fold increase in
PKC activity at 4 minutes, which was significantly inhibited with pretreatment of cells
with PKCα antisense ODNs. A similar result was also seen at 30 minutes, although the
response of PKC to PE had already decreased markedly. The two other antisense PKCα
sequences targeting to 5’ (As2) and coding (As3) sequence of the rat PKCα cDNA had no
effect on the PE induced PKC activity.
5.3.2 Effect of antisense ODN inhibition of PKCα on hypertrophic
phenotype
Expression of two hypertrophic genes, β-MHC and α-SkA, were measured to assess
whether treatment of myocytes with PKCα antisense oligonucleotide was sufficient to
influence cardiomyocyte hypertrophy. Treatment with PE for 48 hours induced a 1.6-fold
increase in α-SkA and a 2.1-fold increase in β-MHC mRNA levels (Fig 9). Inhibition of
PKCα was sufficient to inhibit α-SkA gene expression by 30%, whereas β-MHC gene
expression was not affected (Fig. 9).
Effect of PKCα inhibition on PE induced calcineurin activity was determined next to
examine whether PKCα is an upstream regulator of calcineurin. PE evoked a 4-fold
induction in calcineurin activity (1.18 ± 0.12 vs 0.29 ± 0.08 µM PO4 released).
Transfecting the cells with PKCα antisense ODNs had no significant effect on
calcineurin activity.
To examine the role of alpha subunit of PKC on PE induced ERK activity, an assay
measuring transfer of phosphate by active ERK was used. While PE induced a rapid
activation of ERK, antisense PKCα treatment did not have a significant effect on ERK
58
activity at 4 minutes or at 30 minutes. Interestingly, at 24 hours the PE induced increase
in ERK activity was partially inhibited by antisense PKCα ODN treatment.
β-MHC mRNA
α-SkA mRNA
5
*
1,5
*
#
1
Fold vs control
Fold vs control
2
**
4
**
3
**
2
**
1
0,5
PE
Veh
-
Veh
+
As1
+
Scr
+
**
PE
Veh
-
Veh
+
As1
+
**
Scr
+
Fig. 9. Effect of antisense PKCα ODN treatment on α-SkA and β-MHC mRNA levels in
cultured neonatal rat ventricular myocytes. Veh, vehicle; Scr, scrambled ODN. Results are
expressed as mean ± SEM. *P < 0.05, **P < 0.01 compared with untreated control cells,
#P < 0.05 compared with scrambled ODN treated cells.
PE and ET-1 have been shown to increase intracellular calcium concentration transients
in cultured neonatal cardiac myocytes (Eble et al. 1998, Furukawa et al. 1992). Using
single-cell imaging of Fluo-3-loaded (10 µM) cultured cardiac myocytes, PE was found
to induce a significant increase in intracellular calcium levels (218 ± 37%, p < 0.001).
Antisense PKCα ODN treatment had no effect on PE or ET-induced calcium
concentration transients or spontaneous myocyte beating rate.
To study the effect of PKCα inhibition on protein synthesis, incorporation of [3H]
leucine incorporation into myocytes and total protein content of the myocytes were
measured. Treatment of myocytes with PE for 24 hours induced a 2.7-fold increase in
leucine incorporation and a 1.5-fold increase in total protein content. Treatment of cells
with antisense PKCα ODNs had no effect on PE induced increase in protein synthesis or
total protein content. In agreement with previous studies, ET-1 induced 2.5-fold increase
in [3H] leucine incorporation, which was not affected by antisense PKCα ODN treatment.
Neither did antisense PKCα ODN treatment have any effect on PE or ET-1 induced
increase in myocyte size or myofilament organization.
5.3.3 Effect of antisense PKCα treatment on ET-1 and PE induced
natriuretic peptide secretion and gene expression
Treatment of myocytes with antisense PKCα ODNs had no effect on basal ANP or BNP
secretion. PE (100 µM) treatment induced an up to 32-fold and 15-fold increase in ANP
and BNP release, associated with a 3.1-fold and 1.8-fold increase in ANP and BNP
59
mRNA levels, respectively. Inhibition of PKCα with antisense ODNs significantly
inhibited PE induced ANP release (Fig. 10), whereas it had no effect on PE induced BNP
release (Fig. 10). Antisense treatment had no effect on PE induced increase in ANP or
BNP mRNA levels.
To further characterize the significance of PKCα on hypertrophic signaling, effects of
PKCα inhibition on ET-1 induced natriuretic peptide release and gene expression were
also studied. ET-1 (100 nM) led to an up to 3-fold and 10-fold increase of ANP and BNP
secretion, respectively. Interestingly, antisense inhibition of PKCα resulted in a marked
decrease in both ET-1 induced ANP and BNP secretion (Fig. 10). Similarly to PE
stimulus, inhibition of PKCα had no effect on ET induced increase in ANP or BNP
mRNA levels.
60
A
B
ir-ANP
ir-BNP
**
35
18
30
Scr
**
25
20
15
10
##
##
**
**
**
14
Fold vs basal
Fold vs basal
**
16
12
As1
Scr
10
8
6
As1
4
5
2
0
15
30
45
60
0
15
C
45
60
D
ir-ANP
ir-BNP
4
12
**
**
3
**
#
2
#
**
10
Scr
As1
Fold vs basal
Fold vs basal
30
Time (h)
Time (h)
**
**
8
6
#
Scr
##
##
4
As1
2
1
0
15
30
Time (h)
45
60
0
15
30
45
60
Time (h)
Fig. 10. Effect of antisense PKCα (As1) or scrambled (Scr) ODN treatment on PE (A and B)
or ET-1 (C and D) induced ANP and BNP secretion. Results are expressed as mean ± SEM.
*P < 0.01, **P < 0.001 compared with untreated control cells, #P < 0.05, ##P < 0.001
compared with scrambled ODN treated cells.
61
5.4 Regulation of atrial gene expression by MAP kinases (IV)
5.4.1 Effect of MAPK inhibition on stretch induced natriuretic peptide
gene expression and secretion
Adult rat heart was dissected and the left atrial auricle was attached to the end of a
perfusion tube consisting of two lumens (Laine et al. 1994). The auricle was perfused
with modified Krebs buffer for 30 minutes, thereafter the auricle was stretched by raising
the end of the buffer outflow tube, and thereby increasing the pressure inside the auricle.
p38 MAP kinase activity was increased to 3-fold following a 10-minute stretch. After a
30-minute stretch, the p38 MAPK activity had returned close to basal level. Stretch also
induced a time-dependent increment in ERK activity that peaked at 10 minutes.
Immunoblot analysis with anti-non-phosphospecific antibodies for p38 and ERK showed
that comparable amounts of samples were quantitated in these experiments.
Stretching of atrial preparate for 90 minutes induced a 1.9-fold increase in BNP gene
expression. Inhibition of p38 MAP kinase with SB 203580 (5 µM) resulted in a 30%
decrease of both basal and stretch induced BNP gene expression (Fig. 11). Induction of
the BNP gene expression in the stretched atria was accompanied by a 2.9-fold increase in
atrial IR-BNP levels. Administration of p38 MAP kinase inhibitor SB 203580 (5 µM)
resulted in a 40% decrease in BNP secretion. A 90 minute stretch was not sufficient to
induce ANP gene expression, while a 3.2-fold increase was induced in ANP secretion.
Inhibition of p38 with SB 203580 (5 µM) inhibited ANP secretion by 30% at the 8minute time point. Administration of ERK inhibitor PD 98059 (5 µM) had no effect on
ANP and BNP gene expression and secretion in the stretched atria.
BNP mRNA
**
200
% of control
#
150
*
100
50
Ctrl
Stretch
SB
SB+Stretch
Fig. 11. Effect of p38 MAP kinase inhibition on stretch induced increase in BNP mRNA
levels. Results are expressed as mean ± SEM. **P < 0.001, *P < 0.01 compared with vehicle,
#P < 0.05 compared with stretch.
62
5.4.2 Effect of atrial stretch on cardiac gene expression
The increase in atrial stretch resulted in a transient increase in c-fos mRNA levels, which
was not significant at the 90-minute time point (Fig. 12). Inhibition of p38 MAP kinase
by SB 203580 (5 µM) decreased basal c-fos gene expression by 42% and in the stretched
auricle, gene expression was attenuated by 36% (Fig. 12). β-MHC mRNA levels, in turn,
were elevated by 73% in response to increased stretch. p38 inhibition by SB 203580 (5
µM) was sufficient to decrease the stretch induced increase in β-MHC mRNA levels by
60% (Fig. 12). Interestingly, p38 inhibition induced a slight, but not significant increase
in β-MHC mRNA levels in non-stretched preparates (Fig. 12).
c-fos mRNA
150
β-MHC mRNA
**
200
100
**
75
#
50
25
Ctrl
Stretch
SB
SB+Stretch
% of control
% of control
125
#
150
100
#
50
Ctrl
Stretch
SB
SB+Stretch
Fig. 12. Effect of p38 MAP kinase inhibition on c-fos and β-MHC gene expression in perfused
rat atria. Results are expressed as mean ± SEM. **P < 0.001, compared with vehicle,
#P < 0.05 compared with stretch.
5.4.3 Effect of stretch on transcription factors
Transcription factor Elk-1 has been shown to participate in the regulation of GPCR
agonist induced c-fos gene expression (Babu et al. 2000). Therefore, possible
involvement of transcription factor Elk-1 in stretch activated gene expression was also
studied. Stretching of the auricle was found to induce an increase in serine (ser-383)
phosphorylation of Elk-1. The increase in phosphorylation was already detectable after 5
minutes’ stretching and persisted until 15 minutes. Inhibition of p38 MAP kinase by SB
203580 (5 µM) significantly augmented the stretch induced serine phosphorylation of
Elk-1. To examine whether increased serine phosphorylation also resulted in functional
changes in Elk-1, gel mobility shift analysis was used to measure Elk-1 binding on EBS
in rat BNP promoter (Pikkarainen et al. 2003). Similarly to Elk-1 phosphorylation,
stretching of the auricle induced a rapid increase in Elk-1 binding to BNP promoter
persisting until 30 minutes. At the end of the 90-minute experiment, Elk-1 binding had
returned to basal levels. In accordance with Elk-1 phosphorylation, p38 MAP kinase
inhibition with SB 203580 (5 µM) resulted in a significant decrease in Elk-1 binding to
63
BNP promoter. Inhibition of ERK with PD 98059 (5 µM) had no effect on Elk-1 binding
activity. Stretch also induced a 35% increase in HIF-1α mRNA levels, which was totally
abolished by administration of both PD 98059 (5 µM) and SB 203580 (5 µM).
5.4.4 Regulation of BNP gene expression via EBS
Fold activation
(Relative to basal)
To further characterize the significance of Elk-1 binding on the BNP promoter, we
introduced cultured atrial myocytes with a 534 bp BNP promoter construct with Elk-1
binding site mutated at -498 bp. Stretching of the myocytes for 24 hours activated the
proximal rBNP construct, and the activation was inhibited by the mutation of EBS (Fig.
13). Basal rBNP promoter activity was not significantly affected by mutation of EBS.
3,5
3
2,5
2
*
#
1,5
1
EBS mutation
-
+
Control
-
+
Stretch
Fig. 13. Effect of ETS binding site mutation on basal and stretch induced proximal (-534 bp)
rat BNP promoter activity. Results are expressed as mean ± SEM. *P < 0.001, compared with
basal -534BNP promoter activity, #P < 0.05 compared with stretch induced -534BNP
promoter activity.
6 Discussion
6.1 Role of GATA4 in the regulation of myocyte hypertrophy
GATA4 transcription factor has been shown to play an important role in cardiac
development and in the regulation of cardiac-specific gene expression (Orkin 1995,
Charron & Nemer 1999). The six GATA members can be divided into two subfamilies;
GATA1, -2 and -3, which are expressed in hematopoietic cells, and GATA4, -5 and -6,
which are expressed in various tissues, including the heart. Binding motifs for GATAfactors have been identified within the promoters of most cardiac-expressed genes.
Adenoviral overexpression of GATA4 enhanced sarcomeric organization, induced an
increase in cell surface area and induced a significant increase in total protein
accumulation in cultured neonatal cardiac myocytes (Liang et al. 2001a). Transgenic
mice with 2.5-fold overexpression of GATA4 in the adult heart demonstrated an increase
in heart to body weight ratio, histological features of cardiomyopathy, and activation of
hypertrophic genes, such as ANP, BNP and α-SkA (Liang et al. 2001a).
It has been recently shown that pressure overload and hypertrophic agonists regulate
gene expression via GATA4-dependent mechanism (Hautala et al. 2001). Hypertrophic
agonists activate GATA4-dependent gene expression by increasing GATA4 binding or
transactivating activity (Liang & Molkentin 2002). To assess whether GATA4 binding to
DNA was sufficient and necessary for the development of hypertrophic phenotype and
hypertrophic gene expression, we employed a novel decoy mechanism using GATA
binding decoy oligonucleotides to block GATA4 DNA binding.
GATA decoy ODNs were transfected to cardiac myocytes with high efficiency
compared to previous studies (I). Gel mobility shift analysis showed that GATA decoy
ODNs efficiently blocked both basal and ET-1 induced GATA4 binding to BNP promoter.
Blockade of GATA4 DNA binding resulted in a significant decrease of basal ANP and
BNP secretion and gene expression. Surprisingly, ET-1 induced secretion and gene
expression of both ANP and BNP remained elevated. Previously, GATA4 depletion by
antisense strategy has been shown to downregulate baseline secretion of ANP to a similar
extent (Charron et al. 1999). The decoy strategy used here not only blocks GATA4
binding to the promoter, but also prevents possible compensation by increased binding
65
activity of GATA5 and GATA6. This is a clear benefit over GATA4 antisense strategy,
since it has been reported that GATA6 mRNA levels were increased in GATA4 deficient
mice as a potential compensatory mechanism (Molkentin et al. 1997, Kuo et al. 1997). To
study further the significance of GATA binding sites within the BNP promoter, a 538 bp
BNP promoter luciferase construct was studied. The results were in accordance with
those revealed by the decoy strategy. Mutation of two proximal GATA binding sites in the
BNP promoter decreased basal BNP transcription activity, but it had no effect on ET-1
induced increase in promoter activity.
ET-1 treated cardiac myocytes exhibited a marked increase in sarcomeric assembly
and protein synthesis, as seen in hypertrophied heart in vivo (Hunter & Chien 1999).
Blockade of GATA4 binding by GATA decoy ODNs was not sufficient to prevent these
changes. Adenoviral overexpression of GATA4 was previously shown to enhance
sarcomeric organization and increase total protein accumulation (Liang et al. 2001a).
This implies that GATA4 may participate in hypertrophic gene expression without direct
binding to a gene promoter, probably by interacting with other transcription factors
present in the nucleus, such as Nkx2.5, NFAT3, FOG-2 and MEF2. Results of the current
study (I) demonstrate that blockade of GATA4 binding to a gene promoter has a specific
effect on basal cardiac natriuretic peptide gene expression, but it does not compromise
ET-1 induced natriuretic peptide gene expression. This suggests that direct interactions of
GATA4 with GATA motifs within cardiac gene promoters are not required for ET-1
induced promoter activity. Other characteristics of hypertrophic phenotype induced by
ET-1, such as sarcomeric organization and increased protein synthesis, do not appear to
require direct interaction of GATA4 with gene promoters, either.
6.2 Distinct roles of MAP kinases in the regulation of myocyte
hypertrophy
MAP kinases, ERK, JNK and p38, have been shown to regulate a broad range of
biological functions in response to extracellular stimuli (Sugden & Clerk 1998b). Each
MAP kinase pathway is a multistep cascade facilitating the amplification of the signal.
On the other hand, MAP kinases are known to cross at multiple levels, leading easily to
misinterpretation when studying the MAP kinases. There is an increasing body of
evidence implicating involvement of GATA4 in the hypertrophic signaling in cardiac
myocytes (Hasegawa et al. 1997, Molkentin et al. 1998, Morimoto et al. 2000, Liang et
al. 2001a). GATA4 transcription factor has been shown to be a target for phosphorylation
by ERK in cardiac myocytes (Morimoto et al. 2000). In fact, GATA4 protein has 7 serine
residues, which are potentially phosphorylated by MAP kinases. The aim of the present
study (II) was to examine the role of MAP kinase signaling in the development of
hypertrophic phenotype induced by ET-1 in cardiac myocytes.
ET-1 induced a rapid activation of ERK and p38 MAP kinase, which were totally
abolished by pretreatment of cells with specific inhibitors, PD 98059 and SB 203580,
respectively (Clerk et al. 1998b). ET-1 induced de novo protein synthesis of neonatal rat
ventricular myocytes was also inhibited by pharmacological blockade of p38 MAP
66
kinase, but not with blockade of ERK signaling. These results are rather different
compared to a study which had previously suggested that p38 inhibition has no effect on
ET-1 induced protein synthesis (Choukroun et al. 1998). Another interesting finding in
the study (II) was the induction of p38 activity by ERK inhibitor PD 98059. Previously, a
high dose of PD 98059 (50 µM) has been shown to increase basal levels of
phosphorylated p38 MAP kinase (Clerk et al. 1998b). In turn, activation of ERK pathway
by constitutive active MEK1 (an upstream kinase of ERK pathway) has been shown to
inhibit p38 MAP kinase activity and p38 induced phosphorylation of TATA-binding
protein (Carter & Hunninghake 2000). The inhibition was suggested to be mediated by
MKP-1, which has been shown to block ET-1 induced activation of the MAP kinases
(Carter & Hunninghake 2000, Bueno et al. 2001). Since ERK is necessary for the
stimulation of MKP-1 mRNA expression, the blockade of ERK for 24 hours in the
present experiments is likely to inhibit MKP-1 expression, and thus result in increased
p38 activity (Haneda et al. 1999). Hypertrophic agonists are known to activate MKP-1,
thereby explaining the lack of additive effect with PD 98059 on ET-1 induced p38
activity. Another mechanism involved may be the substrate specificity of MKP-1, since it
has been shown to preferentially block the activation of p38 MAP kinase (Franklin &
Kraft 1997).
A major finding in this study (II) was the differential regulation of GATA4 binding
activity by MAP kinases. Blockade of ERK pathway led to decreased phosphorylation of
serine residues in GATA4 and decreased basal binding activity, but it had no effect on ET1 induced increase in GATA4 DNA binding. ERK overexpression led to phosphorylation
of the serine residues of GATA4 protein, but it was not sufficient to increase GATA4
binding to BNP gene. Blockade of p38 pathway similarly decreased phosphorylation of
serine residues in GATA4, and in contrast to ERK inhibition, totally abolished ET-1
induced GATA4 binding to BNP gene. It is noticeable that p38 overexpression not only
phosphorylated serine residues in GATA4 protein, but also increased GATA4 binding to
BNP promoter. The p38 MAP kinase was also shown to induce BNP promoter activity in
GATA4 dependent manner, since mutation of two proximal GATA binding sites in the
BNP promoter abolished p38 induced BNP promoter activity. Previously, these GATA
binding sites have been shown to direct cardiac myocyte specific expression of rat BNP
promoter and regulate basal promoter activity (Grepin et al. 1994, Thuerauf et al. 1994).
Our results also revealed that overexpression of ERK or JNK pathway members also
induced BNP promoter activation, but independently of GATA binding.
The present results show that blockade of p38 MAP kinase pathway abolishes
hypertrophic agonist induced GATA4 binding to BNP gene, whereas inhibition of ERK
pathway only disrupts GATA4 binding activity in nonstimulated myocytes. Inhibition of
GATA4 binding also results in functional changes in BNP promoter activity. Taken
together, results of the study (II) demonstrate that activation of p38 MAP kinase is
necessary for hypertrophic agonist induced GATA4 binding to BNP gene and sufficient
for GATA-dependent BNP gene expression.
67
6.3 PKCα signaling in myocyte hypertrophy
There are several previous studies indicating the involvement of PKC in the development
of cardiac hypertrophy (Shubeita et al. 1992, Clerk et al. 1994, Sugden & Clerk 1998b,
Braz et al. 2002), yet there are only limited data concerning the precise roles of different
PKC isozymes in hypertrophic signaling. Translocation of PKC isozymes by pressure
overload has provided indirect evidence of PKC participation in hypertrophic signaling.
The objective of this study (III) was to evaluate the significance of PKCα in
cardiomyocyte hypertrophy. Due to the lack of specific pharmacological inhibitors,
studies concerning the role of different isozymes of PKC in the development of cardiac
hypertrophy have been accomplished by using antisense strategy or dominant negative
constructs (Sugden & Clerk 1998b, Molkentin & Dorn II 2001, Strait, III et al. 2001).
In the current study (III), antisense oligonucleotide targeting 3’-untranslated sequence
on the rat PKCα cDNA was most effective in inhibiting PKCα protein synthesis, which is
similar to findings in previous studies using antisense PKCα ODNs (Benimetskaya et al.
2001, Dean et al. 1994). Other cardiac isozymes of PKC were not affected by antisense
PKCα treatment approving the specificity of the antisense. The PKC activity assay also
revealed a significant decrease in agonist induced PKC activity in PKCα antisense ODN
treated cells. Antisense PKCα treatment had no effect on PKC activity in resting cells,
which is probably due to low basal PKC activity. PKCα protein expression was also
studied by cytochemistry, which showed significantly decreased levels of PKCα in
antisense PKCα treated cardiac myocytes.
Cardiac hypertrophy is associated with an increase in cell size and sarcomeric
organization. In the present (III) study, inhibition of PKCα with antisense ODNs was not
sufficient to prevent PE induced increase in cell size or sarcomeric organization.
Interestingly, antisense PKCα treatment significantly decreased PE induced gene
expression of sarcomeric protein α-SkA. It is therefore intriguing to speculate that a
longer experiment time could well reveal attenuation in sarcomere assembly. In fact, a
recent study using dominant negative form of PKCα showed a marked attenuation in PE
induced increase in sarcomeric organization and cell surface area (Braz et al. 2002).
Contrary to the present results, inhibition of PKCα with a dominant negative adenovirus
also showed a marked decrease in PE induced leucine incorporation. The reason for these
contradicting results may be the longer lasting effect on PKCα protein levels produced by
dominant negative PKCα compared to antisense PKCα treatment.
Treatment of cardiac myocytes with dominant negative PKCα also resulted in a
marked decrease in PE induced ANP staining in cardiac myocytes. In the current study
(III), PE and ET-1 both produced a drastic increase in ANP and BNP secretion and gene
expression. Antisense inhibition of PKCα was sufficient to decrease PE induced ANP
secretion and ET-1 induced secretion of both ANP and BNP. Another interesting finding
was that antisense PKCα treatment had no effect on PE or ET-1 induced increase in ANP
or BNP mRNA levels. Cellular mechanisms regulating natriuretic peptide secretion are
not fully understood, but there is a previos study suggesting that ANP secretion from
atrial myocytes is stimulated both by the increase in calcium levels and activation of PKC
(Suzuki et al. 1992). BNP secretion, in turn, is augmented by the activation of PKC,
rather than the elevation in intracellular calcium levels (Suzuki et al. 1992).
68
Interconnection of PKCα with other signaling pathways was also studied. As
previously reported, we found ERK as a downstream target for PKCα (Sugden & Clerk
1998b, Braz et al. 2002). There are also previous data demonstrating that activation of
calcineurin signaling increases α-SkA mRNA levels. In the current study (III), PE
induced a significant activation of calcineurin, which was not affected by inhibition of
PKCα. Therefore it is more likely that calcineurin is an upstream regulator of PKC, as
also previously suggested (De Windt et al. 2000a).
6.4 MAPK regulation of stretch induced cardiac gene expression
Multiple signaling pathways are involved in the regulation of cardiac gene expression
during hemodynamic load, among them PKC, MAP kinases, calcineurin and intracellular
calcium (Sugden & Clerk 1998b, Hunter & Chien 1999, Nicol et al. 2000, Molkentin &
Dorn II 2001). Cardiomyocyte stretch is known to result in increased activity of several
nuclear effectors leading to transcriptional activation of pattern of genes including
structural proteins, such as α-SkA and β-MHC, and natriuretic peptides, ANP and BNP
(Sadoshima & Izumo 1997). The factors transducing the signal from the cell membrane
to the nucleus are not well characterized. The results of this study (IV) suggest that
stretch induced activation of c-fos, β-MHC and BNP gene expression is regulated by p38
MAP kinase and transcription factor Elk-1 in the perfused atria.
Hemodynamic stress induces activation of ERK and p38 MAP kinase cascades and
their downstream effectors in cardiac myocytes (Liang et al. 2000, Sugden 2001). In the
current study, atrial stretch provoked a rapid activation of ERK and p38 MAP kinase
cascades. Transcription factor Elk-1 represents a known target for MAP kinases. In fact,
there is evidence that Elk-1 is activated by ERK, JNK and p38 MAPK pathways
(Treisman 1996, Yang et al. 2001). Interestingly, each of the promoters, c-fos, β-MHC
and BNP, possesses an ETS-binding domain. Previously, Elk-1 together with SRF has
been shown to regulate c-fos gene expression (Aoyagi & Izumo 1993). In fact, minimal
c-fos promoter with a point mutation at the p62TCF binding site of the SRE failed to
respond to pressure overload (Aoyagi & Izumo 1993). In addition to synergic mechanism
with SRF, ETS domain transcription factors have also been shown to act without adjacent
SRE in the promoter to regulate cardiac gene expression (Gupta et al. 1998). Recently,
p38 MAP kinase induced Elk-1 binding on EBS within the proximal rat BNP promoter
was shown to mediate ET-1 induced BNP gene expression in cultured neonatal rat
ventricular myocytes (Pikkarainen et al. 2003).
In the current study, atrial stretch induced phosphorylation of Elk-1 and increased Elk1 binding activity on the rat BNP promoter. Inhibition of p38 MAP kinase, but not ERK,
decreased stretch induced Elk-1 phosphorylation and Elk-1 DNA binding activity. To
further characterize the significance of Elk-1 binding on the BNP promoter, cultured
atrial myocytes were transfected with a proximal rat BNP promoter construct with the
EBS site mutated. 24-hour cyclic stretch was sufficient to induce a 2.9-fold increase in
the activity of intact BNP promoter. Mutation of EBS at -498 significantly decreased
stretch induced BNP promoter activity. Previously, mechanical load induced BNP gene
69
expression has been shown to converge via ET-1 dependent mechanism in the atria, but
not in the ventricle (Magga et al. 1997). Together with the current findings, this proposes
a novel mechanism regulating atrial BNP gene expression in response to increased load.
6.5 Future perspectives
Findings of the current studies show that multiple signaling pathways are involved in
hemodynamic load induced cardiac gene expression, and further help to define the roles
and interrelationships among the regulatory pathways (Fig. 14). While the methods used
here may prove useful in future studies, certain reservations need to be taken into
consideration. Decoy oligonucleotides, which bind to DNA binding site of a transcription
factor, are useful for studying the significance of a transcription factor binding to target
genes, but still allow the transcription factor to control gene expression via interactions
with other nuclear effectors. Antisense ODNs hybridizing to target mRNA, as well as
decoy ODNs, have short half-lifes and their transfection efficiency may prove
insufficient. Instead, post-transcriptional gene silencing by RNA interference would be
useful for studying the roles of specific genes. Overexpression of MAPK pathway
members could also be enhanced by simultaneous overexpression of a constitutively
active upstream kinase, i.e. coexpression of constitutively active MKK3 with wild type
p38α. However, overexpression of a specific kinase may result in a shortage of scaffold
proteins assembling the kinase pathway, and therefore allow unwanted leaking of the
signal to other kinase pathways. It is to be expected that additional experimental tools,
such as usage of small interfering RNAs and adenoviral vectors expressing different
kinases, will allow for cardiac gene regulation to be more fully explored.
Fig. 14. A schematic view of the results of current studies defining the roles of cellular
signaling elements contributing to hemodynamic load induced cardiac gene expression.
7 Summary and conclusions
Cardiac hypertrophy is seen in both physiological and pathophysiological conditions.
Myocardial infarction, essential hypertension, chronic heart failure and cardiomyopathies
are associated with increased cardiac pressure and/or volume overload. Prolonged
overload leads to pathologic cardiac hypertrophy, which involves changes at the level of
gene transcription, protein synthesis and myofibrillar organization.
There are numerous intracellular signaling events contributing to hypertrophic gene
expression. Direct myocyte stretch and GPCR agonists induce activation of intracellular
signaling cascades, such as PKC and MAP kinase pathways, which ultimately affects
activation of transcription factors, including GATA4 and Elk-1, and induces hypertrophic
growth programme of individual myocytes.
1. GATA4 transcription factor is required for normal cardiac development. However, it
is unknown whether GATA4 is an essential mediator of hypertrophic responses in
the heart. The present study demonstrated that GATA decoy treatment efficiently
blocks ET-1 induced GATA4 binding to BNP gene promoter. GATA decoy
treatment significantly decreased basal ANP and BNP secretion and gene expression,
but it had no effect on ET-1 induced natriuretic peptide secretion or gene expression.
Nor did the depletion of GATA binding have any effect on ET-1 induced protein
synthesis or sarcomeric organization. The present results emphasize that GATA4
DNA binding is not necessary for ET-1 induced development of hypertrophic
phenotype.
2. All three MAPK pathways, ERK, JNK and p38, are activated by ET-1 in cultured
cardiac myocytes. Pharmacologic inhibition of p38 MAP kinase attenuated ET-1
induced GATA4 binding to BNP gene promoter and serine phosphorylation of
GATA4. Inhibition of ERK cascade with MEK1 inhibitor PD 98059 reduced basal
and p38 induced GATA4 binding activity, but it had no significant effect on ET-1
induced GATA4 binding activity. Overexpression of p38 MAPK pathway, but not
ERK or JNK pathways, activated GATA4 binding to BNP gene promoter and
induced rat BNP promoter activity via proximal GATA binding sites. These findings
demonstrate that activation of p38 MAPK is necessary for hypertrophic agonist
induced GATA4 binding to BNP gene and sufficient for GATA-dependent BNP gene
expression.
71
3. ET-1 and PE induce activation of PKCα in cultured cardiac myocytes. Antisense
PKCα treatment reduced PKCα mRNA and protein levels and reduced PE induced
PKC activity. Inhibition of PKCα was sufficient to inhibit α-SkA gene expression,
but had no effect on β-MHC gene expression. Inhibition of PKCα with antisense
ODNs significantly inhibited PE induced ANP release, and ET-1 induced release of
both ANP and BNP. Antisense PKCα treatment had no effect on agonist induced
increase of ANP and BNP mRNA levels or on ET-1 or PE induced protein synthesis
or sarcomeric organization. These results suggest that PKCα isozyme is involved in
hypertrophic signaling in cultured cardiac myocytes, but has a minor role in the
development of hypertrophic phenotype.
4. Isolated atrial preparate was used as a model to study the role of ERK and p38 MAP
kinase in stretch induced cardiac gene expression. p38 MAP kinase, but not ERK,
regulated stretch induced transcription factor Elk-1 phosphorylation and DNA
binding activity. Atrial stretch also induced an increase in HIF-1α mRNA levels,
which was attenuated by inhibition of both ERK and p38 MAPK. Administration of
p38 inhibitor also decreased atrial c-fos, β-MHC and BNP gene expression. Studies
with cultured rat atrial myocytes demonstrated that ETS binding site within the rat
BNP promoter was necessary to confer full BNP promoter activation in response to
increased stretch. These findings demonstrate that stretch induced BNP gene
expression is regulated by p38 MAP kinase mediated activation of transcription
factor Elk-1.
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