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University of Iowa
Iowa Research Online
Theses and Dissertations
2011
The intercalated disc-associated Xin family of
proteins in cardiac development and function
Qinchuan Wang
University of Iowa
Copyright 2011 Qinchuan Wang
This dissertation is available at Iowa Research Online: http://ir.uiowa.edu/etd/2653
Recommended Citation
Wang, Qinchuan. "The intercalated disc-associated Xin family of proteins in cardiac development and function." PhD (Doctor of
Philosophy) thesis, University of Iowa, 2011.
http://ir.uiowa.edu/etd/2653.
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THE INTERCALATED DISC-ASSOCIATED XIN FAMILY OF PROTEINS IN
CARDIAC DEVELOPMENT AND FUNCTION
by
Qinchuan Wang
An Abstract
Of a thesis submitted in partial fulfillment
of the requirements for the Doctor of
Philosophy degree in Biology
in the Graduate College of
The University of Iowa
May 2012
Thesis Supervisor: Professor Jim Jung-Ching Lin
1
ABSTRACT
Intercalated discs (ICDs) are cardiac-specific structures located at the longitudinal
termini of cardiomyocytes. Classically, the functions assigned to ICDs include
mechanical and electrical communication among adjacent cardiomyocytes. More
recently, it has been increasingly realized that ICDs also function in signal transduction
and regulation of the surface expression of ion channels. Accordingly, defects of ICD
components are shown to cause a number of human cardiac diseases and changes of ICDs
are associated with cardiomyopathy, arrhythmias, and heart failure. The expansion of our
knowledge about the development, function and maintenance of ICDs is promoted by
identification, cataloging and characterization of the molecular components of the ICDs.
In this thesis, I characterize a family of Xin repeat-containing proteins, which are striated
muscle-specific and localized to the ICDs in the cardiomyocytes. This thesis provides
novel insights into the mechanism of the maturation, maintenance and functions of ICDs.
Our previous studies showed that the Xin repeat-containing proteins play critical
role in cardiac morphogenesis and cardiac function. Knock down of the Xin gene in
chicken embryos collapses the walls of developing heart chambers and leads to abnormal
cardiac morphogenesis. In mammals, paralogous genes, Xinα and Xinβ, exist. Ablation of
the mouse Xinα (mXinα) does not affect heart development. Instead, the mXinα-deficient
mice show late-onset cardiac hypertrophy and cardiomyopathy with conduction defects.
The ICD structural defects in mXinα-null mice occur postnatally between 1 and 3 months
of age and progressively worsen with age. The mXinα-deficient hearts up-regulate
mXinβ, suggesting a partial compensatory role of mXinβ.
In this thesis, I focus on two questions. First, what are the molecular mechanisms
of mXinα’s functions that account for the observed phenotypes in the mXinα-deficient
hearts? And second, what are the functions of mXinβ? Through biochemical methods and
electron microscopy, I demonstrated that mXinα binds and bundles actin filaments. In
2
addition, a direct interaction between mXinα and the adherens junction protein β-catenin
facilitates mXinα’s interaction with the actin filaments. Based on this in vitro
characterization of mXinα, we proposed that mXinα may act as a direct link between the
adherens junctions and actin cytoskeleton, thus providing an important means to
strengthening the intercellular adhesion at the ICDs. To characterize mXinβ’s roles, I
generated and characterized mXinβ-knockout mice. I showed that complete loss of mXinβ
leads to cardiac morphological defects, diastolic dysfunction and heart failure, which lead
to severe growth retardation and early postnatal lethality. I also showed that mXinβ might
be involved in a number of cell signaling pathways and provide multiple lines of
evidence to support mXinβ’s roles in the maturation of ICDs.
In summary, this thesis provides novel insights into the specialization of the
adherens junctions at the ICDs to withstand the contractile forces, and the molecular
mechanisms for the establishment, maintenance and function of ICDs. The knowledge
gained from the roles of Xin proteins in cardiac development and function will likely
provide new insights for improved therapeutic strategies for human cardiomyopathy,
arrhythmias and heart failure.
Abstract Approved: ____________________________________
Thesis Supervisor
____________________________________
Title and Department
____________________________________
Date
THE INTERCALATED DISC-ASSOCIATED XIN FAMILY OF PROTEINS IN
CARDIAC DEVELOPMENT AND FUNCTION
by
Qinchuan Wang
A thesis submitted in partial fulfillment
of the requirements for the Doctor of
Philosophy degree in Biology
in the Graduate College of
The University of Iowa
May 2012
Thesis Supervisor: Professor Jim Jung-Ching Lin
Graduate College
The University of Iowa
Iowa City, Iowa
CERTIFICATE OF APPROVAL
_______________________
PH.D. THESIS
_______________
This is to certify that the Ph.D. thesis of
Qinchuan Wang
has been approved by the Examining Committee
for the thesis requirement for the Doctor of Philosophy
degree in Biology at the May 2012 graduation.
Thesis Committee: ___________________________________
Jim Jung-Ching Lin, Thesis Supervisor
___________________________________
Peter A. Rubenstein
___________________________________
Diane C. Slusarski
___________________________________
Christopher S. Stipp
___________________________________
Chun-Fang Wu
To my parents, Fuqiong Zhang and Guowen Wang
ii
Nature grants not her favors to those with a cold heart.
Santiago Ramón y Cajal, Advice for a Young Investigator
iii
ACKNOWLEDGMENTS
First and for most, I would like to thank Dr. Jim Jung-Ching Lin for his
mentorship, advice, encouragement and support during my graduate studies. I would also
like to thank my committee members: Drs. Peter Rubenstein, Diane Slusarski,
Christopher Stipp, and Chun-Fang Wu for their time, suggestions and guidance. I also
wish to thank Jenny Lin, who has provided invaluable technical assistance and training
over the years, and also encouraged and supported me to overcome all the hurdles. I am
also very appreciative to Rebecca Reiter, who helped me in learning English, techniques
and proofread my manuscripts and comprehensive exam papers. I also wish to thank the
former Lin lab member, Dr. Da-Zhi Wang for his help and suggestions. I wish to thank
Dr. Ming-Che Shih and Hsiao-Ping Peng for their help and encouragement. I would also
like to thank the Lin lab members including Shaun Grosskurth, Sunju Choi, Elisabeth
Gustafson-Wagner, Robbin Eppinga, Shannon Harlan, and Stephen Chan for their help
and discussion. I wish to thank my parents, Fuqiong Zhang and Guowen Wang for their
understanding in their only child’s pursuit of science overseas. And to my wife Keyu
Chen: thank you for your love, support and patience. Finally, I would like to thank my
son Vincent for cheering me up with his babbles and smiles.
iv
ABSTRACT
Intercalated discs (ICDs) are cardiac-specific structures located at the longitudinal
termini of cardiomyocytes. Classically, the functions assigned to ICDs include
mechanical and electrical communication among adjacent cardiomyocytes. More
recently, it has been increasingly realized that ICDs also function in signal transduction
and regulation of the surface expression of ion channels. Accordingly, defects of ICD
components are shown to cause a number of human cardiac diseases and changes of ICDs
are associated with cardiomyopathy, arrhythmias, and heart failure. The expansion of our
knowledge about the development, function and maintenance of ICDs is promoted by
identification, cataloging and characterization of the molecular components of the ICDs.
In this thesis, I characterize a family of Xin repeat-containing proteins, which are striated
muscle-specific and localized to the ICDs in the cardiomyocytes. This thesis provides
novel insights into the mechanism of the maturation, maintenance and functions of ICDs.
Our previous studies showed that the Xin repeat-containing proteins play critical
role in cardiac morphogenesis and cardiac function. Knock down of the Xin gene in
chicken embryos collapses the walls of developing heart chambers and leads to abnormal
cardiac morphogenesis. In mammals, paralogous genes, Xinα and Xinβ, exist. Ablation of
the mouse Xinα (mXinα) does not affect heart development. Instead, the mXinα-deficient
mice show late-onset cardiac hypertrophy and cardiomyopathy with conduction defects.
The ICD structural defects in mXinα-null mice occur postnatally between 1 and 3 months
of age and progressively worsen with age. The mXinα-deficient hearts up-regulate
mXinβ, suggesting a partial compensatory role of mXinβ.
In this thesis, I focus on two questions. First, what are the molecular mechanisms
of mXinα’s functions that account for the observed phenotypes in the mXinα-deficient
hearts? And second, what are the functions of mXinβ? Through biochemical methods and
electron microscopy, I demonstrated that mXinα binds and bundles actin filaments. In
v
addition, a direct interaction between mXinα and the adherens junction protein β-catenin
facilitates mXinα’s interaction with the actin filaments. Based on this in vitro
characterization of mXinα, we proposed that mXinα may act as a direct link between the
adherens junctions and actin cytoskeleton, thus providing an important means to
strengthening the intercellular adhesion at the ICDs. To characterize mXinβ’s roles, I
generated and characterized mXinβ-knockout mice. I showed that complete loss of mXinβ
leads to cardiac morphological defects, diastolic dysfunction and heart failure, which lead
to severe growth retardation and early postnatal lethality. I also showed that mXinβ might
be involved in a number of cell signaling pathways and provide multiple lines of
evidence to support mXinβ’s roles in the maturation of ICDs.
In summary, this thesis provides novel insights into the specialization of the
adherens junctions at the ICDs to withstand the contractile forces, and the molecular
mechanisms for the establishment, maintenance and function of ICDs. The knowledge
gained from the roles of Xin proteins in cardiac development and function will likely
provide new insights for improved therapeutic strategies for human cardiomyopathy,
arrhythmias and heart failure.
vi
TABLE OF CONTENTS
LIST OF TABLES ...............................................................................................................x
LIST OF FIGURES ........................................................................................................... xi
LIST OF ABREVIATIONS ............................................................................................ xiv
CHAPTER I XIN REPEAT-CONTAINING PROTEINS AND INTERCALATED
DISC STRUCTURE, FUNCTION AND FORMATION ................................1 Introduction.......................................................................................................1 Advances in the anatomy of ICD ..............................................................2 The involvement of ICD in signaling ......................................................14 ICDs are formed postnatally ....................................................................20 Discovery, domain structures, expression, and function of the Xin
repeat-containing protein family.....................................................................24 Discovery of the Xin repeat-containing protein family ...........................24 Domain structures of Xin proteins ..........................................................25 Xin is a striated muscle-restricted gene and a downstream target of
Nkx2.5 and Mef2 .....................................................................................26 Two phases of the Xin up-regulation during development correlate
with chamber/valve formation and postnatal heart growth .....................27 Xin expression is significantly up-regulated in animal models of
cardiac hypertrophy and hypertension.....................................................28 The origin of Xin coincides with the first appearance of true heart
chamber, and mXinβ is phylogenetically closer to ancestral Xin
protein than mXinα ..................................................................................29 mXinα plays important roles for the structure and function of the
postnatal heart ..........................................................................................31 Summary and thesis content ...........................................................................33
CHAPTER II THE INTERCALATED DISC PROTEIN, mXINα, IS CAPABLE
OF INTERACTING WITH β-CATENIN AND BUNDLING ACTIN
FILAMENTS ..................................................................................................42 Preface ............................................................................................................42 Abstract ...........................................................................................................43 Introduction.....................................................................................................44 Materials and Methods ...................................................................................46 Yeast Two-Hybrid Assay and Library Screening ...................................46 Constructions of Plasmids and Purification of Recombinant
Proteins ....................................................................................................48 Co-immunoprecipitation(Co-IP), Pull-down Assay and Western
Blot Analysis ...........................................................................................49 Actin Binding Assay................................................................................49 Cell Culture, DNA Transfection and Fluorescence Microscopy.............50 Electron Microscopy ...............................................................................50 Results.............................................................................................................51 mXinα is Associated with N-cadherin, β-catenin and p120 Catenin
in the Adult Mouse Heart ........................................................................51 vii
mXinα Directly Interacts with β-catenin .................................................52 mXinα Binds and Bundles Actin Filaments ............................................54 The β-catenin-Binding Domain and the C-terminal Half of mXinα
Prevent Ectopically Expressed mXinα from Localizing to Stress
Fibers within C2C12 Myoblasts ..............................................................55 Function of the Xin Repeats in Stress Fiber Localization .......................57 The Presence of β-catenin Enhances mXinα Binding to Actin
Filaments In Vitro ....................................................................................59 Discussion .......................................................................................................60 Model for How mXinα Functions at the Adherens Junction of the
Heart ........................................................................................................60 mXinα Contains a Novel β-catenin-Binding Domain .............................63 mXinα Bundles Actin Filaments .............................................................63 CHAPTER III ESSENTIAL ROLES OF AN INTERCALATED DISC PROTEIN,
mXINβ, IN POSTNATAL HEART GROWTH AND SURVIVAL ..............95 Preface ............................................................................................................95 Abstract ...........................................................................................................96 Introduction.....................................................................................................97 Materials and Methods ...................................................................................99 5’ and 3’-RACE (rapid amplification of cDNA ends) of mXinβ
cDNAs .....................................................................................................99 Construction of mXinβ targeting vector and generation of mXinβnull mice ................................................................................................100 Histological staining, immunofluorescence, and assessment of
ventricular myoarchitecture ...................................................................101 Transmission electron microscopy ........................................................102 Proliferation and apoptosis ....................................................................103 Cardiomyocyte width measurement ......................................................103 Body weight, heart weight and liver weight measurements ..................104 Echocardiography ..................................................................................104 Western blot analysis .............................................................................105 Analysis of immunolocalization of N-cadherin and Cx43 during
postnatal heart development ..................................................................106 Rac1 and RhoA activity assay ...............................................................106 Results...........................................................................................................107 Generation of mXinβ-null mice .............................................................107 All mXinβ-null mice die before weaning...............................................108 Loss of mXinβ leads to severe growth retardation ................................109 Loss of mXinβ results in VSDs, abnormal heart shape and misorganized myocardium ..........................................................................110 Developing mXinβ-null hearts exhibit diastolic dysfunction ................110 The delay in switching off slow skeletal troponin I (ssTnI) also
supports diastolic dysfunction associated with mXinβ-null mice ..........112 Developing mXinβ-null hearts exhibit an increased apoptosis as
well as a decreased proliferation ...........................................................112 mXinβ-null hearts fail to develop mature intercalated discs ..................113 The mXinβ-null hearts increased Stat3 activity but decreased Rac1,
IGF-1R, Akt and Erk1/2 activities.........................................................114 Discussion .....................................................................................................115 How does the intercalated disc mature? ................................................116 Diastolic dysfunction may be responsible for heart failure and
lethality in mXinβ-null mice ..................................................................117 viii
mXinβ regulates postnatal cardiac growth ............................................117 CHAPTER IV mXINβ IS ESSENTIAL FOR THE POSTNATAL FORMATION
OF THE INTERCALATED DISCS .............................................................156 Preface ..........................................................................................................156 Abstract .........................................................................................................156 Introduction...................................................................................................157 Materials and Methods .................................................................................161 Animals..................................................................................................161 Antibodies..............................................................................................161 Quantitative Western blot ......................................................................162 Immunostaining .....................................................................................163 Quantification of confocal images.........................................................164 Subcellular fractionation .......................................................................165 Results...........................................................................................................165 The adherens junction proteins, mXinβ, mXinα and N-cadherin
have unique temporal expression profiles during postnatal
development ..........................................................................................165 mXinβ but not mXinα variants preferentially associates with a
subpopulation of N-cadherin at the forming ICDs ................................170 mXinβ preferentially associates with a subcellular fraction
containing the forming ICDs .................................................................171 ICD defects in mXinβ-/- hearts first appear when mXinβ is
expressed at its peak level in the wild-type hearts ................................172 Desmosomes and gap junctions also fail to be restricted to the
termini of cardiomyocytes in mXinβ-/- hearts .......................................173 Intercellular junction components retain their close spatial
relationship in mXinβ-/- hearts despite being mis-localized..................174 mXinα variants are not essential for the formation of ICDs .................175 Discussion .....................................................................................................176 mXinβ plays important roles in the maturation of ICDs .......................177 Molecular mechanisms of ICD formation and mXinβ’s function in
this process ............................................................................................180 Regulation of the expression and localization of mXinβ ......................184 Defects of the mXinβ-null hearts provide novel insights for ICD
formation in healthy and diseased hearts...............................................185 mXinα in the formation and maintenance of ICDs ...............................186 Conclusion .............................................................................................187 CHAPTER V SUMMARY AND FUTURE DIRECTION .............................................214 Overall summary of thesis research ..............................................................214 Conclusion and future direction....................................................................216 APPENDIX A RED/GREEN DOT PROCESSOR .........................................................219 REFERENCES ................................................................................................................225 ix
LIST OF TABLES
Table 2.1. Computer assisted measurements of cell size and shape in transfected
Chinese Hamster Ovary (CHO) cells........................................................................93 Table 2.2. Scoring the population of transfected CHO cells showing GFP fused to
mXinα and to various deletion mutants associated with stress fibers. .....................94 Table 3.1. Genotypes of progenies of mXinβ+/- intercrosses..........................................153 Table 3.2. Echocardiographic analysis of control (mXinβ+/+ & mXinβ+/-) and
mXinβ-null mice at P3.5 and P12.5.........................................................................154 Table 3.3. Assessment of ventricular myoarchitecture ....................................................155 x
LIST OF FIGURES
Figure 1.1. Diagram of an ICD.. ........................................................................................36 Figure 1.2. Major molecular components of the ICD ........................................................38 Figure 1.3. Domain structures of Xin from chick and mouse. ...........................................40 Figure 2.1. Co-immunoprecipitation (Co-IP) of mXinα and β-catenin from adult
mouse heart and from purified recombinant proteins.. .............................................67 Figure 2.2. Determination of the β-catenin-binding domain on mXinα... .........................68 Figure 2.3. Actin binding of purified recombinant His-mXinα and GST-mXinα.. ...........71 Figure 2.4. SDS-PAGE analysis of actin aggregates by GST-mXinα ...............................73 Figure 2.5. SDS-PAGE analysis of low speed actin co-sedimentation with HismXinα. ......................................................................................................................75 Figure 2.6. Characterization of actin bundles formed by His-mXinα and GSTmXinα. ......................................................................................................................77 Figure 2.7. Immunofluorescence microscopy of transiently transfected C2C12
myoblasts. .................................................................................................................79 Figure 2.8. Immunofluorescence microscopy of CHO cells transfected with GFPfull-length mXinα or various GFP-mXinα deletion constructs. ...............................81 Figure 2.9. Yeast two-hybrid assay to demonstrate the interaction of mXinα with
mXinα-interacting proteins. ......................................................................................83 Figure 2.10. Effect of GST-β-catenin on the binding of His-mXinα to actin
filaments....................................................................................................................85 Figure 2.11. Actin bundle formation was accelerated by the presence of GST-βcatenin.. .....................................................................................................................87 Figure 2.12. Schematic model for how mXinα functions at the adherens junction.. .........89 Figure 2.13. Characterization of actin bundles formed by His-mXinα at different
molar ratios of His-mXinα to actin.. .........................................................................91 Figure 3.1. Genomic structure, mRNA and protein isoforms of mXinβ ..........................119 Figure 3.2. Spatial and temporal expression patterns of mXinβ in mice .........................121 Figure 3.3. Generation of mXinβ-null mice. ....................................................................123 Figure 3.4. Neither persistent truncus arteriosus (PTA) nor patent ductus arteriosus
(PDA) was detected in newborn mXinβ-null mouse heart. .....................................125 xi
Figure 3.5. Loss of mXinβ results in severe growth retardation......................................127 Figure 3.6. Structural analyses of mXinβ+/+ and mXinβ-/- hearts. .................................129 Figure 3.7. Doppler flow spectra recorded from the mitral valvular orifices of P12.5
wild type and mXinβ-null mice. ..............................................................................131 Figure 3.8. Masson’s trichrome-stained heart sections from P11.5 wild type and
mXinβ-null mice demonstrating no apparent cardiac fibrosis in the mXinβ-null
heart. .......................................................................................................................133 Figure 3.9. Western blot analysis on total protein extracts prepared from developing
hearts of each mXinβ genotype with anti-myosin heavy chain (MHC)
antibodies, anti-N-cadherin, anti-β-catenin, anti-p120-catenin and DM1B
anti-β-tubulin. .........................................................................................................135 Figure 3.10. A significant delay in switching off ssTnI in mXinβ-null hearts. ................137 Figure 3.11. Increased apoptosis and decreased proliferation in developing mXinβnull hearts................................................................................................................139 Figure 3.12. Representative heart sections from wild type (A, C) and mXinβ-null
(B, D) mice at P3.5 and P12.5 ................................................................................141 Figure 3.13. Mis-localization of N-cadherin and mXinα as well as structural
alteration in developing intercalated disc of mXinβ-null hearts. ............................143 Figure 3.14. The proportion of N-cadherin and connexin 43 localized to the termini
of developing cardiomyocytes of wild type and mXinβ-null mice. ........................145 Figure 3.15. Increased Stat3 activity and decreased Rac1, IGF-1R, Akt and Erk1/2
activities in mXinβ-null hearts. ...............................................................................147 Figure 3.16. No mis-localization of mXinβ in mXinα-null mouse heart. ........................149 Figure 3.17. Proposed roles of mXinβ in postnatal heart growth ....................................151 Figure 4.1. Temporal expression profiles of mXinβ, mXinα variants and Ncadherin in developing postnatal hearts established by quantitative Western
blot.. ........................................................................................................................188 Figure 4.2. Comparison of the expressions of mXin proteins with that of Ncadherin.. .................................................................................................................190 Figure 4.3. Characterization of the co-localization between mXinβ and N-cadherin
during postnatal heart development. .......................................................................192 Figure 4.4. Characterization of the co-localization between mXinα and N-cadherin
during postnatal heart development.. ......................................................................194 Figure 4.5. Subcellular fractionation provided evidence for the preferential
association of mXinβ with the maturing/matured ICDs. ........................................196 xii
Figure 4.6. Time courses of ICD maturation in the postnatal wild-type and mXinβ-/hearts characterized by N-cadherin localization. ....................................................198 Figure 4.7. Characterization of the distributions of desmosome and gap junctions in
the postnatal wild-type and mXinβ-/- hearts. ..........................................................200 Figure 4.8. Confocal images of double labeled frozen sections demonstrating the
preserved co-localization between N-cadherin and desmoplakin in the mXinβ/- hearts. ..................................................................................................................202 Figure 4.9. Confocal images of double labeled frozen section demonstrating the
preserved association between N-cadherin and connexin 43 in the mXinβ-/hearts. ......................................................................................................................204 Figure 4.10. Quantification of the distances between connexin 43 and N-cadherin
immunofluorescence signal spots. ..........................................................................206 Figure 4.11. Confocal images of double labeled frozen sections from P19.5 wildtype and mXinα-/-:mXinβ-/- hearts..........................................................................208 Figure 4.12. Western blot detection of representative intercellular junction proteins
in P13.5 wild-type (lane 1), mXinα-/- (lane 2), mXinβ-/- (lane 3) and mXinα-/:mXinβ-/- DKO hearts (lane 4). GAPDH was used as loading control...................210 Figure 4.13. Quantitative Western blot demonstrated that mXinβ is significantly
down regulated in mXinα-/- hearts at P3.5 and P7.5 but not at P30.5. ...................212 xiii
LIST OF ABBREVIATIONS
adult cTnT
acTnT
amino acid
aa
angiotensin II
AngII
arrhythmogenic right ventricular cardiomyopathy
ARVC
atrium-pulmonary vein
LA-PV
body weight
BW
bovine serum albumin
BSA
bromodeoxyuridine
BrdU
cardiac troponin I
cTnI
Co-immunoprecipitation
Co-IP
connexin 43
Cx43
double knockout
DKO
ejection fraction
EF
electron microscopy
EM
embryonic cTnT
ecTnT
embryonic day
E
embryonic stem
ES
epithelia-mesenchymal transition
EMT
extracellular-signal-regulated kinase 1/2
Erk1/2
fast skeletal troponin I
fsTnI
fetal bovine serum
FBS
fraction shortening
FS
Glutathione S-transferase
GST
glyceraldehyde 3-phosphate dehydrogenase
GAPDH
glycogen synthase kinase 3β
GSK3β
xiv
growth factor receptor-bound protein 2
Grb2
guanine nucleotide exchange factor
GEF
Hamburger-Hamilton stage
HH
heart weight
HW
insulin-like growth factor 1
IGF-1
intercalated disc
ICD
interventricular septum thickness at diastole
IVSd
interventricular septum thickness at systole
IVSs
Janus kinase 2
Jak2
K+ current
IK
knockout
KO
Kv channel interacting protein 2
KChIP2
liver weight
LW
L-type Ca2+ currents
ICa,L
left ventricle posterior wall thickness at diastole
LVPWd
left ventricle posterior wall thickness at systole
LVPWs
left ventricle internal dimension at diastole
LVIDd
left ventricle internal dimension at systole
LVIDs
left ventricle volume at diastole
LVVd
left ventricle volume at systole
LVVs
mitral valve E-wave (early filling) to A-wave (atrial
contraction/late filling) ratio
E/A
Na+ current
INa
NP-40
Nonidet P-40
N-terminal truncated cTnI
cTnI-ND
nuclear export signal
NES
phosphatidylinositol 3-kinase
PI3K
xv
plakophilin 2
PKP2
postnatal day
P
proline-rich region
PR
protein kinase B
Akt
sodium dodecyl sulfate
SDS
signal transducer and activator of transcription 3
Stat3
slow skeletal troponin I
ssTnI
slow skeletal troponin T
ssTnT
suppressor of cytokine signaling 3
SOCS3
T cell/lymphoid-enhancer factors
Tcf/Lef
transient outward K+ currents
Ito
ventricular septal defects
VSDs
α- and β-myosin heavy chain
α- and β-MHC
α-tropomyosin
α-TM
β-catenin-binding domain
β-catBD
β-galactosidase
β-gal
xvi
1
CHAPTER I
XIN REPEAT-CONTAINING PROTEINS AND INTERCALATED
DISC STRUCTURE, FUNCTION AND FORMATION
Introduction
The intercalated discs (ICDs) are essential structures unique to cardiac muscle
(Forbes and Sperelakis, 1985b; Perriard et al., 2003; Severs, 1990); they enable
mechanical coupling and chemical communication among adjacent cardiomyocytes to
achieve regulated contraction for cardiac function. Recent evidence also points to the
involvement of ICD components in transducing signals important for cardiac remodeling
in either the healthy or diseased state (Garcia-Gras et al., 2006; Li et al., 2006; Noorman
et al., 2009; Rohr, 2007; Severs et al., 2008; Sheikh et al., 2009). The structure of the
ICD and its function, deduced from electron microscopic studies, have been
comprehensively summarized in a seminal review paper by Forbes and Sperelakis
(Forbes and Sperelakis, 1985b). In this classical description, the function of ICDs was
assigned to three types of intercellular junctions: (i) gap junctions are responsible for
electrical and chemical communications between cardiomyocytes; (ii) adherens junctions
(fasciae adhaerentes) connect the myofibrils from neighboring cardiomyocytes, thus
transmitting the contractile force; and (iii) desmosomes (maculae adhaerentes) anchor
the desmin intermediate filament to provide mechanical strength to the ICDs. This classic
view of the structure and function of ICDs has been supported by recent studies that
employ genetic, biochemical, physiological and cell biological approaches in several
animal models and cardiac diseases. In general, mutations or deficiencies in ICD
components give rise to many types of cardiomyopathy, arrhythmias and other fatal heart
diseases (for references see recent reviews (Li and Radice, 2010; Noorman et al., 2009;
Perriard et al., 2003; Severs et al., 2008; Sheikh et al., 2009)). Conversely, progression of
2
cardiac disease to heart failure is generally associated with various degrees of ICD
structural disruption.
Recent surveys from the human protein atlas (HPA) web site, ExPASY protein
binding data and published papers reveal nearly 200 proteins associated with ICDs
(Estigoy et al., 2009); about 40% of them are altered in their expression and/or location in
various cardiac diseases (Estigoy et al., 2009). The discovery of a subcellular domain
termed transitional junction (Bennett et al., 2006) between the ICDs and the myofibrils,
further increase the numbers of ICD-associated proteins. Thus, our inventory of ICD
molecular components is far from complete and the molecular mechanisms by which
these components support normal cardiac function remain to be elucidated. Adding to
this list, in 1996, our lab identified a family of Xin repeat-containing proteins in the heart,
which co-localize with adherens junction proteins to the ICDs and play an important role
in cardiac morphogenesis and function (Grosskurth et al., 2008; Gustafson-Wagner et al.,
2007; Wang et al., 1996; Wang et al., 1999; Wang et al., 2010). In this chapter, I will
discuss recent advances in the anatomy of ICDs and in the functions (signaling) of
adhering junctions (adherens junctions and desmosomes), which will be followed by a
brief review of the formation of ICDs. Then, I will summarize what is known about the
Xin repeat-containing proteins and overview the questions I will address in the following
chapters of this thesis.
Advances in the anatomy of ICD
Using immunogold electron microscopy and immunofluorescence microscopy,
two new structures/domains, area composita and transitional junction, at the ICDs were
recently identified (Bennett et al., 2006; Franke et al., 2006). Furthermore,
characterizations of ICD components with molecular, cellular and genetic approches
revealed intricate connections among the intercellular junctions in the ICD.
3
Area composita (mixed type of junctions) exist in
mammalian ICDs but not in non-mammalian ICDs
The adhering junctions (adherens junctions and desmosomes) of the ICD are
traditionally defined based on their morphological resemblance under transmission
electron microscopes to the corresponding junctions in the epithelial cells. The adherens
junction is characterized by a fuzzy electron dense plaque underlining the plasma
membrane. Actin filaments extending from the myofibril thin filaments apparently insert
into the adherens junctions, suggesting that these junctions are the anchorage sites of the
termini of myofibrils. Adherens junctions of the ICD consist of N-cadherin as the
transmembrane component, whose highly conserved cytoplasmic domain interacts with
β-catenin, plakoglobin (γ-catenin), α-catenin (αE-catenin and αT-catenin), p120-catenin,
vinculin and other actin-binding proteins to link to the actin filaments. Conversely, the
desmosomes are characterized by straighter membranes, intermembrane bridges that form
a line in the middle of the gap between two membranes, and the two-layered intracellular
(cytoplasmic) plaques. Intermediate filaments insert into the cytoplasmic plaques. The
desmosomes in the ICD consist of the desmosomal cadherins (desmoglein 2 and
desmocollin 2) and intracellular linker proteins such as desmoplakin and plakophilin 2
(PKP2). Such strict distinction between adherens junctions and desmosomes in the ICD
has been challenged by recent work from Werner W. Franke and his colleagues with
immunoelectron microscopy and immunofluorescence microscopy on mammalian hearts
from different species (Borrmann et al., 2006; Franke et al., 2006). These studies
demonstrated in the adult mammalian ICDs, but not in the non-mammalian ICDs, that the
molecular composition of the adherens junctions and desmosomes are less exclusive than
those in the epithelial junctions (Pieperhoff and Franke, 2007, 2008). The adherens
junctions of the ICD contain not only the typical components of adherens junctions, but
also the desmosomal cadherins and cytoplasmic plaque proteins. Conversely, the
desmosomes in the ICDs contains not only typical desmosomal proteins, but also N-
4
cadherin, β-catenin and α-catenin. Based on these observations, a new type of
intercellular junction, area composita, was proposed (Figure 1.1). The formation of area
composita by fusing adherens junctions and desmosomes appears to be a late process
both in ontogenesis and in evolution (Pieperhoff and Franke, 2007); the area composita is
only found in cardiomyocytes of maturing and adult mammalian hearts (Franke et al.,
2009). The significance of area composita in mammalian hearts remains to be
determined, however, it may strengthen mechanical coupling among neighboring
cardiomyocytes and may enhance crosstalk among different types of junctions. On the
other hand, the absence of the area composita found in the ICDs of non-mammalian
hearts may advantageously assist in the regeneration of damaged hearts. This possibility
is supported by the recent finding that the mouse heart retains impressive regenerative
capacity at birth but not at one week of age (Porrello et al., 2011). The timing of the loss
of regenerative capacity coincides with the maturation of area composita.
Defective adhering junctions generally lead to gap junction
remodeling
Despite the apparent mixing of the molecular components in different types of
junctions of mammalian ICDs, the morphology of adherens junctions and desmosomes
are nevertheless discernible, and the associated filament systems are clearly defined. The
targeted deletion/disruption of mouse genes encoding ICD-associated and actininteracting proteins such as Ena/VASP, mXinα, non-muscle myosin IIB, αE-catenin or
vinculin, seems to affect the morphologically defined adherens junctions specifically and
spare the desmosomes (Eigenthaler et al., 2003; Gustafson-Wagner et al., 2007; Ma et al.,
2009; Sheikh et al., 2006; Zemljic-Harpf et al., 2007). In addition to alterations in the
expression levels of the components of adherens junctions, most of these hearts from the
above mutant animals exhibit reduced levels of connexin 43 (Cx43) expression and
5
altered localization of Cx43 to the lateral side of the cardiomyocytes (gap junction
remodeling).
Gap junction remodeling is also commonly observed in human patient and animal
model hearts with mutations in desmosomal protein components. Desmoplakin is one of
the major components of desmosomes and is capable of interacting with many other
desmosomal components, including PKP2 and plakoglobin, as well as with desmin
intermediate filaments (Sonnenberg and Liem, 2007) (Figure 1.2). The global or cardiac
restricted deletion of desmoplakin in mice results in embryonic lethality, and mutant
embryos display a severe deficiency of desmosomes (Gallicano et al., 1998; Garcia-Gras
et al., 2006); unfortunately, effects on the structures of adherens junctions and gap
junctions of ICD cannot be examined in these mutant lines. However, cardiac-restricted
desmoplakin heterozygous mice recapitulate the phenotype of human arrhythmogenic
right ventricular cardiomyopathy (ARVC), a major cause of sudden cardiac death,
ventricular tachycardia and heart failure (Garcia-Gras et al., 2006). Gap junction
remodeling has also been observed in human patients with ARVC (Basso et al., 2006;
Delmar and McKenna, 2010) due to desmoplakin mutations (Kaplan et al., 2004a; Yang
et al., 2006), plakoglobin deletion (Naxos disease) (Kaplan et al., 2004b) or PKP2
mutations (Fidler et al., 2009). Similarly, cardiac-restricted overexpression of a
desmoplakin missense mutation (disrupting the binding of desmin) results in changing
the expression and localization of Cx43 as well as widening the gaps of ICDs (Yang et
al., 2006). It is also known that gap junction remodeling can occur in human patients with
ischemic cardiomyopathy, dilated cardiomyopathy and heart failure (Bruce et al., 2008;
Dupont et al., 2001; Kaprielian et al., 1998; Kitamura et al., 2002; Kostin et al., 2004;
Kostin et al., 2003; Severs et al., 2008; Smith et al., 1991; Yamada et al., 2003).
In summary, the defective linkage between adhering junctions (adherens junctions
and desmosomes) and the cytoskeleton (actin and intermediate filaments) affects
formation and maintenance of gap junctions (for a more extensive discussion see
6
(Noorman et al., 2009)). As will be described below, gap junction remodeling was
detected in mouse hearts completely lacking one of the Xin repeat-containing and
adherens junction-associated proteins, mXinα. Whether mXinα can directly or indirectly
interact or associate with components of the gap junctions remains to be determined.
Following is a brief review of what is known about junctional proteins that can co-exist
in more than one junction of the ICD.
Linkers involved in molecular crosstalk among different
junctions in ICDs
Recent studies have revealed that many junctional proteins can co-exist in
different junctions, potentially providing linkers to strengthen the mechanical coupling
and to enhance molecular crosstalk. In addition, many protein components of one type of
junction can associate with protein components of another junction (Figure 1.2).
Junctional proteins shuttled and/or linked among different junctions of the ICD could
potentially play important roles in the formation of area composita and/or in molecular
crosstalk among the ICD junctions. Understanding these associations and interactions
may unveil underlying mechanisms for pathogenesis of many cardiac diseases, such as
cardiomyopathy, arrhythmias and heart failure.
Plakoglobin (γ-catenin)
Plakoglobin was the first known junctional component present in both adherens
junctions and desmosomes of the ICD (Cowin et al., 1986). Plakoglobin-null mice die
between E12 and E16 due to severe heart defects (Bierkamp et al., 1996; Ruiz et al.,
1996). In these mutant mice, typical desmosomes are no longer detectable in the heart but
are still present in the epithelial organs, and the desmosomal cadherin, desmoglein 2,
becomes diffusely distributed. The extended adherens junctions of mutant ICDs contain
desmoplakin, most of which co-localizes with β-catenin, thus prematurely forming a
“mixed type” of adhering junction (Ruiz et al., 1996). Furthermore, similar phenotypes
7
have been observed in mice with targeted deletion of PKP2, another armadillo protein
plaque constituent of desmosomes (Grossmann et al., 2004). These results suggest that
both plakoglobin and PKP2 are not only essential for the formation of cardiac
desmosomes, but also critically involved in the segregation of the two sets of molecules
into desmosomes and adherens junctions. Studies with desmoplakin heterozygous
knockout mice and cardiomyocytes in order to understand the pathogenesis of ARVC
have suggested that desmoplakin deficiency leads to mis-localization of plakoglobin from
the ICD to the nucleus. Plakoglobin has structural and functional similarity to β-catenin,
and is able to compete with β-catenin to suppress the Wnt/β-catenin signaling pathway
through T cell/lymphoid-enhancer factors (Tcf/Lef) (Klymkowsky et al., 1999; Zhurinsky
et al., 2000). Suppression of Wnt/β-catenin signaling could promote adipogenic and
fibrogenic gene expression in cardiomyocytes, leading to adipocytic replacement of
cardiomyocytes, the hallmark of ARVC (Garcia-Gras et al., 2006). These results suggest
that plakoglobin can function as a signaling protein in addition to a linking protein
between cadherins and the cytoskeleton. Recent studies with cardiac-restricted
overexpression and deletion of plakoglobin further support this signaling (crosstalk)
function of plakoglobin (Li et al., 2011; Lombardi et al., 2009).
Plakophilin 2 (PKP2)
Mutations in both plakoglobin and PKP2 have been identified in ARVC patients
(Asimaki et al., 2007; Gerull et al., 2004; Kaplan et al., 2004b; McKoy et al., 2000;
Pieperhoff et al., 2008; van Tintelen et al., 2006); about 70% of familial ARVC is caused
by a PKP2 mutation. PKP2 mediates the assembly of desmosomes by scaffolding a
molecular complex containing PKC (protein kinase C), PKP2 and desmoplakin and the
phosphorylation of desmoplakin by PKC is required for desmoplakin’s incorporation into
nascent desmosomes (Malekar et al., 2010). Using small interference RNA (siRNA)
techniques, it has been shown that inhibition of PKP2 expression in primary cultures of
8
neonatal rat ventricular myocytes leads to progressive loss of area composita-like
structures and undetectable desmoplakin in the residual ICD-like structures (Pieperhoff et
al., 2008). Similar to that observed in PKP2-null cardiomyocytes (Grossmann et al.,
2004), knocking down PKP2 also results in an accumulation of many cytoplasmic
vesicles/aggregates containing desmoplakin, PKP2 and desmoglein 2 (Pieperhoff et al.,
2008). These data suggest that PKP2 is involved in the formation and stabilization of the
area composita. In addition, knocking down PKP2 by siRNA causes gap junction
remodeling (a reduction in Cx43 expression, a decrease in dye coupling between cells,
and a significant redistribution of Cx43), further suggesting that PKP2 meditates intraICD crosstalk (Oxford et al., 2007). Thus, PKP2 acts not only as an “organizer” protein
in the formation and stabilization of the area composita, but also function in the
molecular crosstalk between desmosomes and gap junctions (Li and Radice, 2010; Rohr,
2007). The exact mechanism mediating junctional organization and molecular crosstalk
likely involves the multiple functional domains of PKP2, the only plakophilin isoform
expressed in the heart (Mertens et al., 1996). It is known that PKP2 can bind to a large
number of desmosomal proteins, including desmoplakin, plakoglobin, desmoglein and
desmocollin (Chen et al., 2002). Through these interactions, PKP2 may zip up
desmosomal cadherins and tighten the desmosomal plaque. In addition, PKP2 is capable
of interacting with αT-catenin but not αE-catenin; αT-catenin is a component of the
adherens junction which co-localizes with αE-catenin at the ICD (Goossens et al., 2007).
Through its interaction with αT-catenin, PKP2 could link components of adherens
junctions and desmosomes to form and/or stabilize mixed type adhering junctions (area
composita). Furthermore, PKP2 may mediate crosstalk between adhering junctions and
gap junctions through both protein-protein interactions and transcriptional regulation. For
the protein-protein interactions involving PKP2 and gap junctions, it has been shown by
pull-down and co-immunoprecipitation assays from rat heart lysates that PKP2 and Cx43
coexist in the same macromolecular complex; the head domain of PKP2 appears to be
9
sufficient for this association (Oxford et al., 2007). Through this head domain, PKP2 is
also able to associate with β-catenin, which in turn associates, through ZO-1 (zonula
occludens-1) , with the C-terminus of connexin 43 (Wu et al., 2003). At the transcription
regulation level, association of PKP2 with β-catenin up-regulates the signaling activity of
Wnt/β-catenin/Tcf in an overexpression system (Chen et al., 2002), which in turn may
control the Cx43 gene, a known target of Wnt/β-catenin signaling (Ai et al., 2000; van der
Heyden et al., 1998).
p0071 (also referred to as PKP4, plakophilin 4)
Another junctional protein having dual localization in desmosomes and adherens
junctions on epithelial and endothelial cells is p0071 (Calkins et al., 2003; Hatzfeld et al.,
2003). Although this dual localization of p0071 has not been demonstrated in
cardiomyocytes, p0071 message is detected in mouse hearts (Hatzfeld and Nachtsheim,
1996), and its protein product is localized to ICDs (Borrmann et al., 2006). p0071 (PKP4)
belongs to a member of the p120-catenin subfamily of armadillo related proteins; p120catenin is the prototype of this subfamily that comprises p0071, ARVC protein,
NPRAP/δ-catenin and the more distantly related plakophilins 1-3 (Hatzfeld, 2005).
Structurally different from p120-catenin, p0071 contains a PDZ domain-binding motif at
its C-terminus. The head domain of p0071 interacts with desmocollin and desmoplakin,
whereas the armadillo repeat domain binds to classical cadherins (Calkins et al., 2003;
Hatzfeld et al., 2003). In addition, both head and armadillo repeat domains interact with
plakoglobin (Hatzfeld et al., 2003). Moreover, p0071 and p120-catenin can bind to the
same region of the cytoplasmic tail of VE-cadherin, and thus, p0071 can compete p120catenin off from intercellular junctions (Calkins et al., 2003). Functionally similar to
p120-catenin, p0071 can organize small Rho-GTPase signaling, in particular, increasing
RhoA activity via its interaction with Ect2 (a Rho-GEF) and subsequently regulating cell
10
adhesion, cytokinesis and motility (Keil et al., 2007). However, the exact roles of p0071
in ICD formation, stability and function remain to be determined.
p120-catenin
p120-catenin is an armadillo-repeat protein that directly binds to the
juxtamembrane region of classical cadherins and regulates cadherin-based adhesion, cell
shape determination and migration. Recent evidence suggests that like plakoglobin, p120catenin is another component common to adherens junctions and desmosomes, at least in
epithelial cells. In addition to binding E-cadherin, p120-catenin can associate with
desmoglein 1 and desmoglein 3 when desmosomes are assembled in high Ca2+ medium
but not in low Ca2+ medium (Kanno et al., 2008a). These observations of conditional dual
localization suggest that p120-catenin may play an important role both in the regulation
of desmosome assembly and disassembly, as well as in junctional crosstalk. The region
required for the association of p120-catenin with desmosomes has been identified to
aa#758-773 of desmoglein 3, which is different from the plakoglobin-binding site (Kanno
et al., 2008b). However, results from in vitro pull-down assays and yeast two hybrid
assays suggest that p120-catenin cannot directly interact with desmoglein 3 (Bonne et al.,
2003; Kanno et al., 2008b). Similar to p0071, p120-catenin can induce Rac1 and Cdc42
activation via its interaction with Vav2 (a Rho-GEF) and subsequently regulate cell
adhesion, shape and motility (Noren et al., 2000; Noren et al., 2001). However, this
regulatory role of p120-catenin has not been demonstrated in cardiomyocytes. As will be
described below, the loss of Xin repeat-containing and ICD-associated protein, mXinβ, in
the developing heart impairs N-cadherin clustering during the formation of mature ICDs,
alters the expression and localization of p120-catenin, and significantly reduces Rac1
activity (Wang et al., 2010). How mXin proteins influence p120-catenin and Rac1
remains to be determined. Finally, p120-catenin has been shown to link the adherens
junctions to the minus end of microtubules through PLEKHA7 and Nezha, which is
11
required for the establishment and maintenance of the zonula adherens (Meng et al.,
2008).
ZO-1 (zonula occludens-1)
ZO-1 is a member of the membrane-associated guanylate kinase (MAGUK)
family of proteins and originally discovered in association with the tight junction
(Stevenson et al., 1986). In the heart, ZO-1 is localized in endothelial cells, interstitial
cells and at the ICDs of cardiomyocytes (Barker et al., 2002; Bruce et al., 2008;
Toyofuku et al., 1998). The N-terminal half of ZO-1 contains 3 PDZ domains, a SH3
domain and a catalytically inactive guanylate kinase domain (Itoh et al., 1997; Toyofuku
et al., 1998). Through its second PDZ (PDZ-2) domain, ZO-1 binds to the extreme Cterminus of Cx43 (Giepmans, 2004; Giepmans and Moolenaar, 1998; Toyofuku et al.,
1998). The recombinant N-terminal of ZO-1 can also bind directly to α-catenin, whereas
the C-terminal specifically co-sediments with actin filaments in vitro and localizes to
microfilament bundles in non-muscle cells (Itoh et al., 1997). Therefore, ZO-1 is able to
crosstalk between gap junctions and adherens junctions. Supporting the importance of
crosstalk between gap junctions and adherens junctions, it has been shown that the
associations between adherens junctional proteins and Cx43 are required for the
development/formation of gap junctions in non-muscle cells (Shaw et al., 2007; Wei et
al., 2005) as well as cardiomyocytes (Wu et al., 2003). In addition, as described above,
depletion of the Cx43-associated desmosomal protein, PKP2, by siRNA treatment of
culture cardiomyocytes leads to gap junction remodeling and a decrease in dye coupling
between cells (Oxford et al., 2007; Pieperhoff et al., 2008). Therefore, molecular
crosstalk between adhering junction components and gap junction proteins at the ICD
may account for the underlying mechanisms for gap junction remodeling observed in
many human cardiac diseases and heart failure.
12
In the heart, Cx43-associated ZO-1 may also play a key role in regulating size,
number and distribution of gap junctions (Hunter et al., 2005; Palatinus and Gourdie,
2007). ZO-1 was found to preferentially localize to the periphery of gap junction plaques,
presumably either to inhibit further recruitment of connexons or to favor their removal
from gap junctions on reaching a certain size (Barker et al., 2002; Bruce et al., 2008;
Hunter et al., 2005). Within the ICDs in vivo, only low level co-localization between ZO1 and Cx43 is found, as compared with the relatively high level co-localization between
ZO-1 and N-cadherin (Barker et al., 2002). However, during remodeling of cardiac gap
junctions, such as in the enzymatically isolated cardiomyocytes (Barker et al., 2002) or in
the human failing heart (Bruce et al., 2008), co-localization and interaction between ZO-1
and Cx43 strikingly increase. This increased interaction of Cx43 with ZO-1 could
constrain the growth of gap junctions and contribute to reduction in the Cx43 levels
observed in the human failing heart (Bruce et al., 2008; Dupont et al., 2001; Kostin et al.,
2003; Severs et al., 2008). Further support of this hypothesis comes from studies of both
in vitro cell systems (Hunter et al., 2005) and in vivo hearts of mice expressing Cterminally truncated Cx43 (K258stop/KO) (Maass et al., 2007). Specific disruption of the
interaction between ZO-1 and Cx43 leads to increased size, decreased number, and
altered localization of gap junction plaques. Thus, both ZO-1 and the C-terminal domain
of Cx43 are involved in regulating the organization of Cx43 plaques. As will be
described below, mXinα-null mouse hearts display gap junction remodeling (GustafsonWagner et al., 2007). Like ZO-1, mXinα has been shown to be a β-catenin-binding and
actin-binding protein located at ICDs (Choi et al., 2007). It would be of interest to
investigate whether mXinα and ZO-1 may cooperatively be involved in the assembly and
maintenance of gap junction plaques in the heart.
13
Transitional junction
A recent study has also advanced our understanding of how sarcomeres are
connected to the ICD (Figure 1.1). It has long been noticed that in addition to the
intercellular junction covered membrane, the ICD membrane also contains regions free of
gap junctions, adherens junctions and desmosomes (Forbes and Sperelakis, 1985a).
Bennett and coworkers observed that these regions are mainly located at the apex of the
membrane interdigitations and are associated with spectrins (Bennett et al., 2006). It has
been shown that this junction-free region of the ICD membrane is at the level where the
Z-disc of the last sarcomere would have been located, if the last sarcomere formed a Zdisc close to the ICD. Interestingly, some Z-disc proteins such as α-actinin, titin, ZASP
are identified in this region. In addition, although the thin filaments extend from the
sarcomeres into the ICD seamlessly, the ICD actin seems to be β-actin instead of the
sarcomeric α-actin (Balasubramanian et al., 2010). This isoform switch appears at the Zdisc-like region, where non-muscle myosin IIB (Ma et al., 2009) and NRAP (Manisastry
et al., 2009; Zhang et al., 2001) are also found; this specialized Z-disc-like structure was
thus defined as the transitional junction (Bennett et al., 2006). Bennett et al.’s work
explains how the last sarcomere retains regular organization even though its thin
filaments insert into the highly convoluted ICD. It also suggests that new sarcomeres can
be added onto the end of the myofibril in the convoluted region of ICDs without
disturbing the overall organization of the myofibrils. Indeed, addition of new sarcomeres
in the ICD was recently observed in cardiomyocytes whose myofibrils are elongating
under volume overload (Yoshida et al., 2010). The ICDs change the organization of their
interdigitation to accommodate the addition of forming sarcomeres without disrupting the
overall organization of the myofibrils, supporting an important role for the ICD in
myofibril formation.
14
The involvement of ICD in signaling
Recent evidence from studies with transgenic overexpressing and knockout
animals clearly points to the involvement of ICD components in transducing signals
important for cardiac remodeling in both physiological and pathological states (see
references in (Bass-Zubek et al., 2009; Garcia-Gras et al., 2006; Li et al., 2006; Li et al.,
2011; Lombardi et al., 2009; Noorman et al., 2009; Rohr, 2007; Severs et al., 2008)).
Here, we only briefly discuss signaling relevant to β-catenin in the heart, because the βcatenin-binding domain is present in Xin repeat-containing proteins (Choi et al., 2007). βcatenin is a multifunctional protein and plays a central role in regulating both canonical
Wnt (Wnt/β-catenin) signaling and cadherin-mediated (cadherin/β-catenin) signaling in
many cell types and tissues (Kwiatkowski et al., 2007; Nelson and Nusse, 2004; PerezMoreno and Fuchs, 2006). The interplay between these two signaling pathways has been
shown to be crucial in the process of epithelia-mesenchymal transition (EMT), which
occurs not only in normal embryonic development, but also in tumor formation and
metastasis (Heuberger and Birchmeier, 2010). Both Wnt/β-catenin and N-cadherinmediated signaling pathways likely operate in the postnatal and adult hearts, and a faulty
component of these pathways could result in cardiac hypertrophy and cardiomyopathy
(Chen et al., 2006; Garcia-Gras et al., 2006; Hirschy et al., 2010; Li et al., 2006; Li et al.,
2011; Lombardi et al., 2009). In the presence of canonical Wnt signaling, cytoplasmic βcatenin is stabilized and enters the nucleus, where it interacts with T-cell factors (TCFs),
such as lymphoid enhancer factor 1 (Lef1), to regulate gene expression. In the absence of
Wnt signaling, cytoplasmic β-catenin is targeted for destruction by the APC, axin, and
GSK3β complex that phosphorylates β-catenin and directs it to a destruction pathway
(Nelson and Nusse, 2004). β-catenin is known to bind N-cadherin at ICDs to regulate Ncadherin-mediated adhesion. Therefore, canonical Wnt and N-cadherin-mediated
signaling pathways potentially compete for the same pool of β-catenin.
15
N-cadherin/β-catenin signaling in the heart
It has been shown that either too much or too little of N-cadherin in the heart
leads to dilated cardiomyopathy, suggesting that delicate signaling through N-cadherin is
required for normal adult heart function. Transgenic mice over-expressing N-cadherin in
the heart develop cardiomyopathy, whereas ectopic expression of E-cadherin in the heart
leads to a much more severe cardiomyopathy (Ferreira-Cornwell et al., 2002). Ectopic
expression of E-cadherin in the heart would interfere with the N-cadherin-mediated
signal and result in a more severe cardiomyopathy. Conditional deletion of N-cadherin in
the adult heart leads to a complete dissolution of ICD structure and a significant decrease
in the gap junction protein, Cx43 (Kostetskii et al., 2005). Consequently, N-cadherindeficient mice exhibit dilated cardiomyopathy, impaired cardiac function, ventricular
arrhythmias and sudden death (Li et al., 2008; Li et al., 2005). These results suggest that
the N-cadherin-mediated adhesion and signaling pathway are essential for structural
integrity and function of the heart.
The most characterized cellular signals involving cadherin/catenin complexes are
those generated locally upon cadherin-cadherin engagement during cell-cell contact
formation. In non-cardiomyocytes, the small GTPases (Rho, Rac and Cdc42) have been
shown to transduce such local signals to control cell adhesion, survival, shape and
motility (Arulanandam et al., 2009; Raptis et al., 2009; Watanabe et al., 2009). Following
the engagement, juxtamembrane domain of cadherin interacts with p120-catenin, which
can activate Rac1 and Cdc42, by binding to Vav2, a guanine nucleotide exchange factor
(GEF) for these GTPases (Noren et al., 2000). In addition to initiating cellular signals
during contact formation, cadherin/catenin complexes in established junctions are also
involved in mediating signal transduction. The adherens junctions are recognized as a
sensor for mechanical forces and transduce signals that influence the actin cytoskeleton.
Such mechanical signal transduction appears to rely on the proteins linking adherens
junctions to the actin cytoskeleton (le Duc et al., 2010; Yonemura et al., 2010), and likely
16
involves the small GTPases (Smutny and Yap, 2010). Interestingly, we have shown that
mXinα is capable of interacting not only with β-catenin but also with p120-catenin (Choi
et al., 2007). Furthermore, the mXinβ-null heart showed a significant decrease in active
Rac1, a failure to form mature ICD and a misaligned myocardium (Wang et al., 2010).
These results together suggest an involvement of Xin repeat-containing proteins in the Ncadherin-mediated signaling pathway.
Wnt/β-catenin in the heart
The role for Wnt/β-catenin signaling in cardiac development has been intensively
studied in a variety of organisms, although controversy remains. During early cardiac
development, Wnt/β-catenin signaling appears to have developmental stage-specific
biphasic effects on cardiogenesis (Naito et al., 2006; Ueno et al., 2007). Activation of
Wnt/β-catenin signaling before gastrulation promotes mesoderm formation and
cardiogenesis, whereas signaling during and after gastrulation inhibits cardiomyocyte
differentiation by opposing bone morphogenetic protein signaling (Marvin et al., 2001;
Tzahor and Lassar, 2001). However, the hypothesis that Wnt actively inhibits
cardiogenesis is still too simple. Recent studies using both gain and loss of Wnt/β-catenin
function have shown that Wnt/β-catenin pathway acts cooperatively with FGF and BMP
signaling to promote expansion of the second heart field progenitors (Ai et al., 2007;
Cohen et al., 2007; Klaus et al., 2007), which contribute to outflow tract and right
ventricle (Buckingham et al., 2005; Srivastava, 2006). In postnatal and adult hearts, the
importance of Wnt/β-catenin signaling for cardiac remodeling at physiological and
pathological conditions has also been intensively addressed. Activation of β-catenin in
cultured rat neonatal cardiomyocytes was found to be not only sufficient but also
necessary to induce cardiomyocyte hypertrophy (Force et al., 2007; Haq et al., 2003). In
vivo studies using inducible cardiac-specific knockout or transgenic mice to modulate the
expression levels of Wnt/β-catenin signaling components or their mutants have further
17
confirmed that stabilization of β-catenin or activation of Wnt signaling is required for
both physiological and pathological cardiac hypertrophy (Chen et al., 2006; Malekar et
al., 2010; Qu et al., 2007; van de Schans et al., 2007). However, conflicting results have
also been reported (Baurand et al., 2007). The precise reason for such discrepancy is
unknown but may reflect the pleiotropic effects of Wnt signaling depending on the
experimental conditions. Recently, studies with conditional transgenic mice expressing
either no β-catenin or stabilized β-catenin generated by using a ventricle-specific driver
(MLC2v-Cre) have revealed that mice lacking β-catenin in the adult ventricles do not
have an overt phenotype (Hirschy et al., 2010), due to an up-regulation of plakoglobin, as
suggested previously (Zhou et al., 2007). In contrast, mice expressing stabilized β-catenin
develop cardiac hypertrophy and dilated cardiomyopathy at 2 months of age, and do not
survive beyond 5 months (Hirschy et al., 2010). Furthermore, the stabilized β-catenin
was only found at the ICDs but never detected in the nucleus (Hirschy et al., 2010).
These results suggest that β-catenin’s role in nucleus may be of little significance in the
healthy adult heart, and that similar to N-cadherin, too much β-catenin at ICD may
critically affect the N-cadherin/β-catenin signaling and subsequently lead to dilated
cardiomyopathy. It should be noted that increased β-catenin levels were also detected in
the hypertrophic hearts from human cardiomyopathy patients and from spontaneously δsarcoglycan-deficient hamsters (Masuelli et al., 2003). The accumulation of β-catenin at
ICD, but not in nucleus, is accompanied by an increased Wnt5a (a noncanonical Wnt)
expression, a decrease in GSK3β expression and a differential expression of APC
isoforms. The existence of multiple Wnt signaling pathways in the heart has added
another level of complexity to Wnt signaling related to cardiac remodeling.
18
Interplay between Wnt/β-catenin signaling and adhering
junction-mediated signaling
Down-regulation of Wnt/β-catenin signaling by nuclear plakoglobin detected in
ARVC hearts might be part of the molecular mechanism for the pathogenesis of ARVC
(Garcia-Gras et al., 2006; Lombardi et al., 2009). Adult mice heterozygous for the
conditional deletion of desmoplakin in the heart recapitulate phenotype of ARVC
(Garcia-Gras et al., 2006). Apparently, the desmoplakin deficiency leads to an impaired
desmosome assembly, which could free plakoglobin from the desmosomes and increase
its nuclear localization in cardiomyocytes. Plakoglobin is known to be able to compete
with β-catenin at multiple cellular levels with a net negative effect on the Wnt/β-catenin
signal pathway (Klymkowsky et al., 1999; Zhurinsky et al., 2000). Thus, increasing
plakoglobin nuclear localization in desmoplakin heterozygous mice should suppress
Wnt/β-catenin signaling, which in turn would promote adipogenesis, fibrogenesis and
apoptosis (Chen et al., 2001; Longo et al., 2002; Ross et al., 2000), the characteristic
hallmarks of human ARVC. This mechanism has been further supported by two recent
studies with transgenic mice over-expressing plakoglobin in cardiomyocytes as well as
mice with conditional knockout of plakoglobin in cardiomyocytes, respectively. Overexpressed plakoglobin translocates to nucleus and suppresses Wnt/β-catenin signaling.
The association of plakoglobin, instead of β-catenin, with Tcf712 increases the
expressions of Wnt5b and BMP7, which promote adipogenesis, and decreases the
expression of connective tissue growth factor, which is an inhibitor of adipogenesis
(Lombardi et al., 2009). The adipocytes in mouse and human ARVC hearts were
identified to originate from the second heart field progenitors, accounting for a
predominant involvement of right ventricle in human ARVC (Lombardi et al., 2009). On
the other hand, Wnt/β-catenin signaling was activated in the hearts of mice with inducible
cardiac-restricted plakoglobin deletion (Li et al., 2011). Upon deletion of plakoglobin,
expression levels of Wnt/β-catenin target genes, such as c-Myc and c-Fos, were increased
19
significantly. Stabilization of β-catenin following the loss of plakoglobin may be due to
activation of Akt and inhibition of GSK3 (Li et al., 2011), which could affect β-catenin
phosphorylation state/stability and promote cardiac hypertrophy. Interestingly, stabilized
β-catenin in plakoglobin-null heart became associated with Tcf4, a transcription factor
primarily binding to plakoglobin. These results are consistent with the idea that β-catenin
directly competes with plakoglobin for Tcf4 binding. The mutant mice exhibited
progressive loss of cardiomyocytes, extensive inflammation, fibrosis, altered desmosome
structure and cardiac dysfunction similar to ARVC patients. However, in contrast to the
desmoplakin conditional knockout hearts, neither adipocyte replacement nor lipid droplet
accumulation was observed in the conditional plakoglobin knockout hearts, suggesting
that plakoglobin itself might be required for the full spectrum of ARVC phenotype.
Thus, based on the mouse models with various manipulations of the components
of the adhering junctions (adherens junctions and desmosomes), it seems that the
molecular mechanism of AVRC consists of both nuclear and desmosomal signaling
pathways. In one mechanism, elevated plakoglobin in the nuclei alters Wnt/β-catenin
signaling, which appears to be critical for the manifestation of ARVC phenotype such as
apoptosis, fibrosis and adipogenesis. On the other hand, signals generated by the
desmosomes likely play a role in the pathology of the ARVC, because down-regulation
of Wnt/β-catenin signaling by simply conditional deletion of β-catenin in the postnatal
heart does not lead to ARVC. The specific involvement of signals from the desmosomes
in ARVC is further supported by the lack of ARVC phenotype in mice with conditional
deletion of adherens junction components, such as N-cadherin (Kostetskii et al., 2005),
αE-catenin (Sheikh et al., 2006), mXinα (Gustafson-Wagner et al., 2007) and β-catenin
(Chen et al., 2006; Hirschy et al., 2010; Zhou et al., 2007). These mice exhibit dilated
cardiomyopathy with neither myocyte loss nor inflammation, which is different from the
conditional plakoglobin knockout mice (Li et al., 2011) and other animal models of
ARVC (Lombardi et al., 2009; Pilichou et al., 2009; Yang et al., 2006). These
20
differences suggest that a different signaling may transduce through adherens junctions
versus desmosomes.
ICD influences ion channel surface expression
As a functional unit, the ICD also plays important roles in organizing and/or
regulating surface ion channels and receptors. Previous studies have shown that the poreforming α-subunit, Nav1.5, of the voltage-gated sodium channel is preferentially
localized to the ICD (Cohen, 1996; Kucera et al., 2002; Maier et al., 2004; Maier et al.,
2002; Mohler et al., 2004b). This population of sodium channel complexes is composed
of Nav1.5, tyrosine-phosphorylated β1 subunit, and ankyrin G in close association with
both N-cadherin and Cx43 (Malhotra et al., 2004; Meadows and Isom, 2005). A recent
study has shown that Nav1.5 can be pulled down by the head domain of PKP2 from heart
lysates, suggesting that PKP2 participates in the same molecular complex at ICDs (Sato
et al., 2011). Knockdown of PKP2 expression in cultured cardiomyocytes by siRNA
leads to a decrease in peak current density, changes in the current kinetics (inactivation
and recovery from inactivation), and a slower velocity of action potential propagation
(Sato et al., 2009). Collaborating with Dr. Cheng-I Lin’s group, we also presented
evidence that ICD-associated mXinα protein influences surface expression of transient
outward potassium current (ITO) through its ability to interact with Kv channel interacting
protein 2 (KChIP2) (Chan et al., 2011), an auxiliary subunit of ITO, and filamin, an actincrosslinking protein. Taken together, these results further suggest a link among all 4
components of the ICD: desmosomes, adherens junctions, gap junctions and the voltagegated channel complex. It seems relevant to consider the ICD as an overall functional
unit when seeking to understand the pathogenesis of cardiac diseases.
ICDs are formed postnatally
The importance of ICDs for the structure and function of the heart leads to
considerable interest in the formation of ICDs. The adult ventricular cardiomyocytes are
21
rod-shaped cells. A structural and functional segregation exists between the parts of the
plasma membrane that are either parallel or perpendicular to the long axis of the cells.
The membrane that is parallel to the long axis (lateral membrane) associates with the
extracellular matrix through costameres. On the other hand, the membrane that is
perpendicular to the long axis (terminal membrane) forms the highly specialized ICDs
that mediate cell-cell communications (Perriard et al., 2003). Early observations with
transmission electron microscope had hinted that formation of ICD is a late event of
cardiac development. With electron microscopy, Legato found that in neonatal dog
hearts, cardiomyocytes were tightly packed together and extensively contacted with each
other, contrasting with the limited cell-cell contacts at the ICDs in adult hearts (Legato,
1979). Forbes and Sperelakis noticed that the ICDs of the postnatal day 2 (P2) mouse
hearts were more primitive than adult ICDs, in that the former had less inter-digitation
and less dense cytoplasmic plaques (Forbes and Sperelakis, 1985b). However, electron
microscopy could only reveal ICD components that are already incorporated into
specialized intercellular junctions; thus the extent of developmental changes during ICD
formation had largely been overlooked until specific antibodies against ICD components
were utilized to study the development of ICD.
Although all the three types of intercellular junctions localized to the adult ICDs
could be identified by electron microscopy in mouse cardiomyocytes at embryonic day
10 (E10) (Forbes and Sperelakis, 1985b), recent studies with specific antibodies
demonstrated that ICDs are formed through a series of events that happen mainly
postnatally. In the mouse and rat, the typical adult ICDs are not completely formed until
P90, while in human, formation and maturation of ICDs continues until age 7 (Angst et
al., 1997; Hirschy et al., 2006; Peters et al., 1994; Pieperhoff and Franke, 2007). The
formation and maturation of ICDs, in essence, is a process of specialization of the
subdomains of the cardiomyocytes’ membrane. During the formation of ICDs,
cardiomyocytes redistribute the adhering cell-cell junctions (adherens junctions and
22
desmosomes) and gap junctions to the terminal ends, and through poorly understood
processes, further recruit an extensive panoply of junctional, channel, signaling and
auxiliary proteins to the ICDs.
Redistribution of Junctional components during ICD
formation
Adherens Junctions: The leading role of adherens junctions in the hierarchy of
establishment and maintenance of different intercellular junctions has been demonstrated
in various in vitro and in vivo systems, including cardiomyocytes (Eppenberger and
Zuppinger, 1999; Hertig et al., 1996a; Hertig et al., 1996b; Kostetskii et al., 2005). Thus
the developmental changes of adherens junctions are particularly important when
studying ICD formation. Indeed, the components of adherens junctions are extensively
redistributed during embryonic and postnatal development in the heart. Consistent with
the tight packing and extensive contacting of cardiomyocytes in the embryonic hearts
(Legato, 1979), immunofluorescence staining localizes N-cadherin to the cardiomyocyte
surface almost homogeneously at E10.5 (Sinn et al., 2002). Later during embryonic
development, N-cadherin staining becomes heterogeneous and the cardiomyocyte surface
is characterized by bright spots interspaced by weakly and diffusely stained area (Sinn et
al., 2002). These bright spots likely represent N-cadherin clusters (and their intracellular
partners) involved in strong homophilic interaction with opposing cells. On the other
hand, the adherens junctional components not incorporated in such bright spots may
represent N-cadherin molecules that are yet to be clustered and engaged in strong cellcell adhesion. Consistent with this idea, adherens junctions identifiable in TEM
micrographs only occupy a small fraction of the cell-cell contacting interface (Legato,
1979), despite the extensive staining of N-cadherin on the surface of cardiomyocytes
during these embryonic stages.
23
The extensive coexistence of brightly stained spots and more diffusely distributed
N-cadherin signal can be found on virtually the entire surface of cardiomyocytes until
P3.5 (Angst et al., 1997; Hirschy et al., 2006; Pieperhoff and Franke, 2007; Sinn et al.,
2002). The trend for N-cadherin to become heterogeneously distributed continues after
P3.5, leading to the eventual loss of N-cadherin staining at lateral surface of
cardiomyocytes and the restriction of N-cadherin to the ICDs at the termini of
cardiomyocytes.
Desmosomes: The dependence of desmosomes on the adherens junctions for their
formation and maintenance has been well documented. Consistent with this, the time
course of incorporation of desmosomal components into the ICDs seems to follow that of
adherens junctions (Angst et al., 1997). In addition, the establishment of area composita
and thus the almost perfect co-localization between adherens junctions and desmosomes
by immunostaining seems to be a prolonged event that lasts from embryonic to postnatal
stages. Different proteins seem to mix together at different developmental stages. In the
mice, the intracellular components of adherens junctions and desmosomes amalgamate
first during embryonic development whereas the transmembrane proteins N-cadherin and
desmoglein-2 are still increasing their level of co-localization even at 3 weeks after birth.
(Pieperhoff and Franke, 2007).
Gap junctions: The developmental redistribution of gap junctions is one of the
first phenomena noticed by researchers demonstrating the late formation of ICDs (Peters
et al., 1994). The incorporation of gap junctions into the cell termini seems to happen
much later than the adherens junctions and desmosomes, and this phenomenon is shared
by different mammalian species examined so far. In the mouse, at 3 weeks postnatally,
prominent Cx43 staining is seen as large puncta located at both the ICD and the lateral
surface (Angst et al., 1997). Quantitative analysis of the distribution of adherens
junctions and gap junctions also supports the delayed incorporation of gap junctions to
the termini of cardiomyocytes (Angst et al., 1997; Peters et al., 1994).
24
Discovery, domain structures, expression, and function of
the Xin repeat-containing protein family
Discovery of the Xin repeat-containing protein family
Prior to the availability of genome-wide microarray and functional genomics, our
lab used differential mRNA display screening in conjunction with whole-mount in situ
hybridization to clone novel genes that are temporally and spatially expressed during
cardiac morphogenesis (Wang et al., 1996). Cardiac cushion formation and valvuloseptal
morphogenesis are essential for a four-chambered heart. These processes involve
inductive interaction between myocardium and endocardium as well as epithelialmesenchymal transformation (EMT), which occur temporally in chicken embryos
between Hamburger and Hamilton (HH) stage 15 and 21 and spatially at the future
atrioventricular (AV) canal and future outflow tract of the linear heart tube (Butcher and
Markwald, 2007; Eisenberg and Markwald, 1995; Markwald et al., 2010). Therefore, we
performed differential display cloning on the total RNAs prepared from AV canal region
of stage 15 and 21 chicken hearts. Whole-mount in situ hybridization was used as a
secondary screening method to confirm the temporal and spatial expression patterns of
isolated genes. From this screen a novel gene, 21C, among others was identified. Later,
we used antisense oligonucleotide treatment of culture chicken embryos to show that this
21C plays a very important function in cardiac morphogenesis and looping (Wang et al.,
1999). Subsequently, we cloned mouse homologs of 21C and identified it being a
downstream target of Mef2C and Nkx2.5 (Wang et al., 1999). Because of its critical role
for normal heart development but not because of its strong cardiac expression as wrongly
cited in Otten et al.(Otten et al., 2010), we then called this gene as Xin (a Chinese word
meaning heart). Xin encodes a modular protein that contains a N-terminal 16-amino acid
repeating unit with a consensus sequence of
GDV(K/Q/R/S)XX(R/K/T)WLFET(Q/R/K/T)PLD (Lin et al., 2005; Pacholsky et al.,
25
2004; Wang et al., 1999). Since the initial discovery of chicken Xin (cXin), two
homologous genes, each containing a Xin repeat region, have been identified in
mammals: mXinα and mXinβ (also called Myomaxin) in mouse (Gustafson-Wagner et al.,
2007; Huang et al., 2006; Sinn et al., 2002; Wang et al., 1999; Wang et al., 2010) as well
as hXinα (also called cardiomyopathy associated 1, CMYA1, or Xin actin-binding repeat
containing 1, XIRP1) and hXinβ (also called CMYA3 or XIRP2) in humans (Lin et al.,
2005; Pacholsky et al., 2004; van der Ven et al., 2006).
Domain structures of Xin proteins
The Xin gene encodes a striated muscle-specific protein containing a region with
15~28 Xin (16-aa) repeats. The Xin repeat defines a new class of actin binding domain
and a minimum of 3 repeats is required to bind actin filaments (Pacholsky et al., 2004). In
chapter II, I will show that the mXinα not only binds but also bundles actin filaments
(Choi et al., 2007). The pink box shown in Figure 1.3 represents the Xin repeat region
found in all Xin proteins from chick and mouse. Within the Xin repeat region, there is a
highly conserved β-catenin-binding domain (β-catBD, indicated by a light green box),
which has been mapped previously on mXinα (Gustafson-Wagner et al., 2007). Similar to
hXinα (van der Ven et al., 2006), mXinα undergoes an unusual intraexonic splicing of
exon 2 to generate two variants differing in its C-terminus (Gustafson-Wagner et al.,
2007). The larger variant is termed mXinα-a, which contains a region homologous to
filamin c-binding motif (red box in Figure 1.3) identified in hXinα (van der Ven et al.,
2006). However, this filamin c-binding motif is not found in cXin and mXinβ
(Grosskurth et al., 2008). Another feature of all Xin repeat-containing proteins is the
existence of multiple proline-rich (PR) regions. The highly conserved PR1 sequence (E/V
BD in Figure 1.3) at the N-terminus has been shown to bind to Mena/VASP proteins
(Grosskurth et al., 2008; van der Ven et al., 2006). Interestingly, the sequences
downstream of the Mena/VASP-binding domain are highly conserved among all Xin
26
proteins (indicated by yellow box in Figure 1.3) and homologous to Myb DNA-binding
domain, despite that the function of this putative DNA binding domain (DBD) is
unknown. The roles of the other PR regions including PR2, PR3 and PR distributed at the
C-terminus of the protein remain unclear.
In mXinβ, alternative splicing of the primary transcript leads to an inclusion or
exclusion of exon 8 and results in two protein variants differing in its very C-terminus
(Wang et al., 2010). The significance of this difference remains to be determined. The
large variant is called mXinβ-a. Both mXinβ and mXinβ-a also possess consensus
sequences for nuclear export signal (NES), nuclear localization signal (NLS) and
ATP/GTP-binding domain (ATP_GTP_A loop) (Wang et al., 2010), however, the
functions of these domains are still unknown.
Xin is a striated muscle-restricted gene and a downstream
target of Nkx2.5 and Mef2
Multiple tissue Northern blot analyses revealed that cXin (9.0kb), mXinα (5.8kb)
and mXinβ (12kb) messages were detected only in the heart and skeletal muscle (Huang
et al., 2006; Lin et al., 2005; Wang et al., 1999). Occasionally, a low level of Xin
expression could be detected in the pulmonary vein of lung tissue, which may represent
the associated cardiomyocytes in this tissue. Whole-mount in situ hybridization revealed
that expression patterns of cXin and mXinα in developing heart and somites (Wang et al.,
1999) are very similar to that of Nkx2.5 and Mef2C (Edmondson et al., 1994; Lints et al.,
1993). In an anterior medial mesoendoderm explant system, the induction of cXin
expression by BMP-2 followed activation of Nkx2.5 and Mef2C, but preceded expression
of the myosin heavy chain (Wang et al., 1999). Similar effects on cardiac looping
morphogenesis observed in embryos after Nkx2.5 (Lyons et al., 1995) or Mef2C (Lin et
al., 1997) deletion or cXin antisense treatment (Wang et al., 1999) further suggest that
cXin participates in a BMP-Nkx2.5-Mef2C pathway to regulate cardiac morphogenesis.
27
Either Nkx2.5 or Mef2C alone is able to trans-activate the expression of luciferase
reporter gene driven by mXinα promoter in non-muscle cells. These results together with
drastically down-regulated mXinα messages in Nkx2.5-null and Mef2C-null embryos (Lin
et al., 2005) suggest that mXinα also participates in a Nkx2.5-Mef2C pathway to control
cardiac differentiation and morphogenesis in mouse. Moreover, mXinβ has been shown
to be a direct target of Mef2A (Huang et al., 2006) and to function downstream of
angiotensin II (AngII) signaling to modulate pathological cardiac remodeling (McCalmon
et al., 2010). Therefore, the Xin repeat-containing protein family plays an important role
in cardiac development and function through the Nkx2.5-Mef2 pathway.
Two phases of the Xin up-regulation during development
correlate with chamber/valve formation and postnatal heart
growth
During embryogenesis, cXin transcript is first detected at HH stage 8 in the paired
lateral plate mesoderm that forms the primordium of the heart (Wang et al., 1999). At
stage 9, cXin expression increases substantially in the heart forming fields, which migrate
anteriorly and ventrally toward the midline of the embryo. At stage 10, cXin is expressed
exclusively in the linear heart tube. Cardiac specific expression continues until stage 15,
when somite expression begins to be detected. Skeletal and cardiac muscle-restricted
expression of cXin continues throughout development and into adulthood. The relative
expression level of cXin/GAPDH determined from Northern blot results of developing
hearts reveals that two major phases of cXin up-regulation exist at stage (st.)16-25 and
post-hatch day (D)12-14 (unpublished data), respectively. These two peaks of cXin upregulation appears to coincide with the timing for chamber/valve formation and postnatal
heart growth (Butcher and Markwald, 2007), suggesting a role of cXin in normal cardiac
morphogenesis. Supporting this role, knocking down the cXin by antisense
28
oligonucleotide collapses the wall of heart chambers, leading to abnormal cardiac
morphology (Wang et al., 1999).
mXin protein was first detected in the developing linear heart tube of E8.0 mice
by whole-mount immunofluorescence microscopy with antibody recognizing both mXinα
and mXinβ (Sinn et al., 2002). At this stage, one to seven somites are present but contain
no mXin protein. At E10 (HH st.17 in chick), mXin is expressed throughout the
myocardium but not endocardium of the truncus arteriosus, common atrial chamber and
ventricle, and meanwhile, mXin expression begins to be detected within the myotome of
the first two rostral most somites (Sinn et al., 2002). As development progresses to E13.5
(HH st.30), staining detects strong mXin expression throughout the myocardium of the
ventricle and atria. Co-localization of mXin with both β-catenin and N-cadherin is
observed to the cell periphery but not the nucleus of early mouse embryonic hearts (Sinn
et al., 2002). In chapters III and IV, I will show detailed analysis of the postnatal
expression profile of mXin proteins and provide evidence to support the important roles
of mXinβ for the postnatal development of the heart.
Xin expression is significantly up-regulated in animal
models of cardiac hypertrophy and hypertension
In response to abnormal stresses, such as hypertension, pressure overload,
endocrine disorders and myocardial infarction, adult cardiomyocytes undergo
pathological hypertrophy. This hypertrophy can be a compensatory mechanism that helps
to preserve pump function in pathological conditions. Frequently, this hypertrophy
progresses to dilated cardiomyopathy. Using pressure overload-induced hypertrophy by
thoracic aortic banding (Hill et al., 2000; Rockman et al., 1993; Rockman et al., 1991),
we detected up-regulation of both mXinα and mXinβ messages in the banded hearts, as
compared with sham-operated control (unpublished data). In addition,
immunofluorescence microscopy showed that banded hypertrophic hearts had thickening
29
ICDs containing much more mXin and N-cadherin proteins (unpublished data),
suggesting that mXin may play an important role in modulating hypertrophy/stress
responses in adult heart. The up-regulation of mXinβ has been also observed in the
hypertensive hearts induced only by Ang II infusion (within 6 hours) but not by salt,
suggesting that its up-regulation is due to Ang II-induced myocardial damages and not to
blood pressure elevation per se (Duka et al., 2006). Thus, the up-regulation of mXinβ
appears to be one of the earliest molecular events triggered by Ang II. It is likely that
mXinα up-regulation would be also observed in this Ang II-induced
hypertension/myocardial damage model, because both mXinα and mXinβ are
transcriptional targets of MEF2 (Huang et al., 2006; Lin et al., 2005; Wang et al., 1999).
It has been recently, shown that mXinβ hypomorphic mice with 80% reduction in mXinβ
message results in cardiac hypertrophy (McCalmon et al., 2010). Hearts from these
hypomorphic mice display less myocardial damage when exposed to Ang II (McCalmon
et al., 2010). These results suggest that mXinβ functioning downstream of Ang II
signaling can modulate cardiac function in health and disease.
The origin of Xin coincides with the first appearance of true
heart chamber, and mXinβ is phylogenetically closer to
ancestral Xin protein than mXinα
A phylogenetic analysis has been performed with 40 vertebrate Xins to elucidate
the evolutionary relationship between Xin proteins and to identify the origin of Xin
(Grosskurth et al., 2008). Multiple sequence alignment (Rambaut, 1995; Thompson et al.,
1994) of the Xin repeats from vertebrates was analyzed in maximum likelihood and
Bayesian analyses (Abascal et al., 2005; Guindon and Gascuel, 2003; Stamatakis et al.,
2005). The constructed evolutionary tree replicates the phylogeny of taxa with the
mammal, other land vertebrates and teleost phylum-level groups. Clearly, the whole
genome duplication which occurred early in evolution produces Xinα and Xinβ proteins
30
(Grosskurth et al., 2008). The additional gene duplication of Xinβ was detected in
teleosts. BLAST searches only detect Xin in the chordates but not in other organisms
such as Saccharomyces cerevisiae, Candida albicans, Arabidopsis thaliana,
Dictyostelium discoideum, Caenorhabditis elegans, Anopheles gambiae (mosquito) and
Drosophila melanogaster. Further analyses identified no Xin repeat-containing proteins
in the Urochordate tunicate (Ciona savignyi) or the Cephalochordate amphioxus
(Branchiostoma floridae). A Xin protein, defined as a protein containing Xin repeating
units, is thus first identified in the Craniate lamprey (evolved about 550 million years
ago). Importantly, both the Urochodate tunicate and the Cephalochordate amphioxus
have a heart with only single layer of contracting mesoderm or contracting vessel coupled
with incomplete endothelial cell layer, whereas the Craniate lamprey has a true
chambered heart with complete endothelial and myocardial layers. Thus, the origin of
Xin proteins coincides with a critical evolutionary modification of the heart, namely, the
origin of true chambers. This finding is consistent with the chamber genesis role of Xin
repeat-containing proteins in vertebrates.
In the avian lineage there is a loss of Xinβ, however, its sole Xin protein still
retains 27 Xin repeats (compared to 28 repeats in the ancestral lamprey Xin) in order to
carry out essential functions of Xin (Grosskurth et al., 2008). Therefore, chicken embryos
treated by antisense oligonucleotide targeting cXin showed defects in cardiac looping and
morphogenesis (Wang et al., 1999). In the mammalian lineage, Xinα contains a reduced
number of Xin repeating units with 17 repeats in opossum and 15 in placental mammals.
This strongly suggests that there was selective pressure within the mammalian lineage
that resulted in the reduction of Xin repeats. Because mXinα-null mice are viable but
develop cardiac defects in adulthood (Gustafson-Wagner et al., 2007) and mXinβ-null
mice die postnatally with chamber defects (to be discussed in chapter III) (Wang et al.,
2010), it is likely that mammalian Xinαs evolved quickly to form many unique traits for
neofunction in adult heart, whereas all mammalian Xinβs are highly conserved with the
31
ancestral lamprey Xin and retain its function in embryonic development (Grosskurth et
al., 2008). Evolutionary study has also identified a putative DNA-binding domain
conserved in the N-terminus of all Xins, in addition to a highly conserved β-catenin
binding domain within the Xin repeat region (Grosskurth et al., 2008). In the C-terminus,
Xinαs and Xinβs are more divergent relative to each other but each isoform from
mammals shows a high degree of within isoform sequence identity (Grosskurth et al.,
2008). These results suggest different but conserved functions for mammalian Xinα and
Xinβ.
mXinα plays important roles for the structure and function
of the postnatal heart
The indispensable role of the cXin in the morphogenesis of chicken heart led us to
study the mammalian Xins. We first generated and characterized mXinα knockout mice
because mXinα is the first known mouse Xin gene (Wang et al., 1999). The mXinα-null
mice are viable and fertile and show no structural abnormalities in the heart at young age;
however, they progressively develop cardiac hypertrophy and cardiomyopathy with
conduction defects. Characterization of the mXinα-/- hearts revealed important roles of
mXinα in the maintenance of the structural integrity of ICDs in the adult heart
(Gustafson-Wagner et al., 2007). We and our collaborators also showed that mXinα plays
important roles in the surface expression of ion channels; thus mXinα is also involved in
the electrophysiology of the heart.
mXin proteins are co-localized with N-cadherin and Cx43 in myocardium of
mouse (Sinn et al., 2002). Loss of mXinα causes a decrease in the expression level of βcatenin, N-cadherin, and desmoplakin in the adult hearts, weakening cardiomyocyte
adhesion and compromising the integrity of the intercalated discs. Consequently Cx43 in
mXinα-null hearts are mis-localized, which may lead to cardiac gap junction remodeling
and conduction defects (Gustafson-Wagner et al., 2007).
32
Studies have revealed more a detailed mechanism by which mXinα affects the
electrophysiology of the heart. Whole-cell patch-clamp recordings on ventricular
myocytes obtained from 10~20-week-old mice revealed that mXinα-null mice have an
increased inward Na+ current (INa), a reduced transient outward K+ currents (Ito), a weaker
L-type Ca2+ currents (ICa,L) and a smaller inward rectifier K+ current (IK1) densities (Chan
et al., 2011; Cheng et al., 2005; Lai et al., 2007). In addition, the amplitude of
intracellular Ca2+ transient decreases significantly in mXinα-null myocytes prepared from
ventricles and left atrium pulmonary vein (LA-PV) (Chan et al., 2011). Optical mapping
analyses revealed that conduction velocity is significantly slower in the mXinα-null than
in the wild-type ventricles and LA-PV (Lai et al., 2007; Lai et al., 2008). These data
suggest the mXinα plays an important role in regulation of channel activity in both
ventricular and atrial myocytes.
mXinα interacts with different proteins that regulate the surface expression of ion
channels, which may account for the altered electrophysiological properties of mXinαnull cardiomyocytes. Prominently, yeast two-hybrid interaction assays revealed that
mXinα interacts with Kv channel interacting protein 2 (KChIP2), an auxiliary subunit of
Ito channel. It is also known that KChIP2 interacts with Kv4.2, the pore-forming αsubunit of Ito channel, and maintains the surface expression of Ito (Guo et al., 2002;
Nerbonne and Kass, 2005). Consistently, the loss of mXinα decreases the expression of
KChIP2 protein and KChIP2 message, leading to significant reduction in surface
expression of Ito current (Chan et al., 2011). Furthermore mXinα has been shown to bind
to filamins and actin filaments (Choi et al., 2007; van der Ven et al., 2006); thus, mXinα
affects the Ito current density by interacting with and stabilizing the KChIP2 and filamin
proteins.
The role of mXinα in regulating ion channel surface expression and function may
be analogous to that of the actin-binding protein ankyrins. The ankyrins interact with
actin filaments and spectrin, and together with their associated proteins form a membrane
33
cytoskeleton, which plays an important role in targeting of ion channels, transporters and
cell adhesion molecules to specialized compartments within the plasma membrane.
Mutations affecting association between ankyrins and ion channels alter functions of ion
channels and result in cardiac arrhythmia. For example, human SCN5A missense
mutation defective in binding to ankyrin-G leads to reductions of INa at T-tubes and ICDs
and results in Brugada syndrome (Mohler et al., 2004a). mXinα interacts with actin
filaments and maintains the structure of ICDs, thus, the reduction of INa current density in
mXinα-null mice may result from altered cytoskeletal structure of the ICDs.
Reduction of channel activity may induce cardiac electrical remodeling. Our
previous studies using optical mapping (Lai et al., 2007) found that the mXinα-deficient
mice had a hypertrophied ventricular myocardium with reduced conduction velocity.
Similarly, the conduction velocity also reduced in LA-PVs from mXinα-null hearts
(29 ± 3 cm/s) compared to wild-type hearts (52 ± 3 cm/s) (Lai et al., 2008). The latter
study also found that the mXinα-null LA-PV have a larger area of conduction block than
the control (defined as the area with a conduction velocity ≤ 10 cm/s; (Eijsbouts et al.,
2004)). Finally, when we used isoproterenol to activate β–adrenoceptor, the mXinα-null
LA-PV are less responsive to the adrenergic activation as compared to wild-type LA-PV.
It is known that Cx43 knockout and loss of N-cadherin decreased the degree of
coupling and conduction characteristics. Thus in mXinα-null mice, decreased cardiac
conduction velocity may result from lower expression of N-cadherin and Cx43 proteins.
Summary and thesis content
Since the first definitive description of ICDs was made in the 1950s with electron
microscopy, our knowledge about this cardiac-specific structure has advanced
significantly. In this chapter, I have reviewed the structure, function and formation of
ICDs. Clearly, the great advancement of our understanding of ICDs is mainly a result of
the discoveries and characterizations of the protein components of the ICDs. In this
34
context, I reviewed the discovery and characterization a novel family of ICD-localized
proteins, the Xin repeat-containing proteins. Our study on the evolution showed a
coincidence of the emergences of Xin genes and true chambered hearts, implicating
important roles of the Xin family for the development and function of the heart. Indeed,
the importance of the Xin proteins in cardiac morphogenesis is supported by the
disruption of heart development by anti-sense oligos against the cXin. Our lab has further
generated and characterized mice that lack one of the mouse Xin genes, mXinα. The
cardiac phenotypes of mXinα-null mice during adulthood strongly support the important
and unique functions of mXinα. However, the fact that mXinα-null animals do not show
lethal phenotype like that observed in the anti-sense oligo-treated chicken embryos also
indicates that the evolutionarily more conserved mXinβ may largely compensate for the
loss of mXinα.
The above discoveries are the foundation on which I built my thesis work. In the
following chapters, I will first introduce my effort in characterizing the molecular
properties of mXinα (charter II). In chapter II, the direct interaction between mXinα and
β-catenin is demonstrated by multiple approaches. In the meantime, chapter II also shows
that mXinα not only binds but also bundles actin filaments, and β-catenin facilitates
mXinα’s interaction with the actin filaments. Based on these observations, a model
depicting the role of mXinα as a direct link between the adherens junctions and the actin
cytoskeleton was proposed. Chapter II further provides a molecular understanding of the
phenotypes observed in the mXinα-deficient mice. In chapter III, I tested our hypothesis
that besides the common functions shared by mXinα and mXinβ, mXinβ plays important
and unique roles for the development and function of the heart. I will describe my work
in generation and characterization of mXinβ-null mice. Consistent with the idea that
mXinβ may have the evolutionarily more conserved roles of the Xin family of proteins,
mXinβ-null animals show neonatal lethality, and cardiac morphological as well as
functional defects. Evidence presented in the chapter III also indicates that mXinβ might
35
be an important scaffolding protein that mediates adherens junction signaling and other
cell signaling pathways. In addition, observations presented in chapter III imply the
indispensible role of mXinβ in the formation of ICDs. In chapter IV, the role of mXinβ
for the formation of ICDs will be further studied. I will provide detailed description of the
formation of ICDs and demonstrate the indispensable roles of mXinβ in this process. In
the last chapter (chapter IV), I will summarize this thesis and propose questions to be
addressed in the future.
36
Figure 1.1. Diagram of an adult mammalian ICD. The adhering junctions (adheres
junctions and desmosomes) intermix to form the area composita. Both actin filaments and
intermediate filaments insert into the area composita. The transitional junctions are
located at the level of the apexes of the membrane folds and mediate the transition
between the thin filaments and the actin filaments in the ICD. The question mark
indicates that the organization of the actin filaments in the ICDs is not well understood.
SR, sarcoplasmic reticulum.
37
38
Figure 1.2. Major molecular components of the ICDs. Proteins that may shuttle between
different intercellular junctions are shown. ZO-1 interacts with Cx43, αE-catenin and βcatenin and thus it may provide a link between the gap junctions and the adherens
junctions. Plakoglobin is known to interact with both N-cadherin and desmosomal
cadherins (desmocolin 2 & desmoglein 2) and thus it may play an important role in the
association between adherens junctions and the desmosomes. αT-catenin may be a direct
link between the adherens junctions and desmosomes through its interaction with the βcatenin and plakophilin 2. p120-catenin is a critical component of the adherens junctions
and it has been shown to associate with the desmosomes. However, the mechanism of
p120-catenin’s association with the desmosomes is not clear; thus such association is not
shown. Plakophilin 2 links the desmosomes and the gap junctions together. P0071 is not
shown because its interaction partner in the ICDs has not been defined.
39
40
Figure 1.3. Domain structures of Xin from chick and mouse. Amino acid residue numbers
are labeled above on each Xin proteins. Through alternate splicing, both mXinα and
mXinβ genes encode two protein variants, which differ only in the very C-terminal
sequences. The actin-binding motifs are contained in the Xin repeat region (indicated by
pink box), within which a conserved β-catenin binding domain (β-catBD, indicated by
light green box) is located. Other binding domains defined on large variant (hXinα-a) of
human Xinα include filamin c-binding region (indicated by red box in mXinα-a) and
Mena/VASP-binding domain (PR1, E/V BD, indicated by purple box in all Xin proteins).
The functions of other proline-rich regions (PR, PR2, PR3) and other consensus
sequences such as DNA-binding domain (DBD), nuclear export signal (NES), nuclear
localization signal (NLS) and ATP/GTP binding domain (ATP_GTP_A loop) remains
unknown.
41
42
CHAPTER II
THE INTERCALATED DISC PROTEIN, mXINα, IS CAPABLE OF
INTERACTING WITH β-CATENIN AND BUNDLING ACTIN
FILAMENTS
Preface
The following chapter includes data prepared for a manuscript by Sunju Choi#,
Elisabeth A. Gustafson-Wagner#, Qinchuan Wang#, Shannon M. Harlan, Haley W. Sinn,
Jenny L-C. Lin, and Jim J-C. Lin (# these authors contributed equally), which has been
published in the Journal of Biological Chemistry (The Journal of Biological Chemistry,
2007; 282: 36024-36036). In this study, we address the molecular mechanisms
underlying the function of mXinα. Specifically, we demonstrate that mXinα directly
interacts with the adherens junction component, β-catenin, and mapped this β-catenininteraction domain to a region within the Xin repeats. We also demonstrate that mXinα
not only binds to, but also bundles actin filaments, and importantly, β-catenin facilitates
these activities of mXinα. From the data presented in the paper, we propose a model for
the mechanism of the functions of mXinα in the heart.
My direct contributions to this paper involved the following: 1) demonstrating the
ability of mXinα to bundle actin filaments by low speed actin co-sedimentation
experiments as well as negative staining and electron microscopy (Figures 2.4, 2.5, 2.6
and 2.13), ; 2) demonstrating that β-catenin facilitates the actin bundling activity of
mXinα through time-course electron microscope observation of the formation of actin
bundles by mXinα in the presence or absence of β-catenin (Figure 2.11). My observations
are instrumental for our understanding of the molecular mechanism of mXinα’s function
in the heart. Based on my observations and the data contributed by Sunju Choi and
Elisabeth A. Gustafson-Wagner, we proposed that mXinα may present in an autoinhibited status and an open (active) status for actin binding and bundling, and β-catenin
43
at the adherens junctions of the intercalated discs promotes mXinα to shift into the open
status for interacting with actin.
Abstract
Targeted deletion of mXinα results in cardiac hypertrophy and cardiomyopathy
with conduction defects (Gustafson-Wagner et al., 2007). To understand the underlying
mechanisms leading to such cardiac defects, the functional domains of mXinα and its
interacting proteins were investigated. Interaction studies using co-immunoprecipitation,
pull-down and yeast two-hybrid assays revealed that mXinα directly interacts with βcatenin. The β-catenin-binding site on mXinα was mapped to amino acid #535-636,
which overlaps with the known actin-binding domains composed of the Xin repeats. The
overlapping nature of these domains provides insight into the molecular mechanism for
mXinα localization and function. Purified recombinant GST- or His-tagged mXinα
proteins are capable of binding and bundling actin filaments, as determined by cosedimentation and electron microscopic studies. The binding to actin was saturated at an
approximate stoichiometry of 9 actin monomers to one mXinα. A stronger interaction
was observed between mXinα C-terminal deletion and actin as compared to the
interaction between full-length mXinα and actin. Furthermore, expression of GFP-fused
to mXinα C-terminal deletion in cultured cells showed greater stress fiber localization
compared to expressed GFP-mXinα. These results suggest a model whereby the Cterminus of mXinα may prevent the full-length molecule from binding to actin, until the
β-catenin binding domain is occupied by β-catenin. The binding of mXinα to β-catenin at
the adherens junction would then facilitate actin binding. In support of this model, we
found that the actin binding and bundling activity of mXinα was enhanced in the
presence of β-catenin.
44
Introduction
The striated muscle-specific Xin genes encode proteins containing several prolinerich regions, a highly conserved sequence homologous to the Myb-A and Myb-B DNA
binding domain, and a region with 15~28 16-amino acid (aa) repeating units (called the
Xin repeats) (Lin et al., 2005; Pacholsky et al., 2004; Wang et al., 1999). In the mouse,
two Xin genes, mXinα and mXinβ, exist, whereas only one cXin gene is found in the
chick. The expression of both cXin and mXinα is regulated by the muscle transcription
factor, MEF2C, and the homeodomain transcription factor, Nkx2.5 (Lin et al., 2005;
Wang et al., 1999). The expression of mXinβ (also termed myomaxin) is under the
control of MEF2A (Huang et al., 2006). Treatment of chick embryos with cXin antisense
oligonucleotides results in abnormal cardiac morphogenesis and a disruption in cardiac
looping, suggesting that Xin plays an essential role in cardiac development (Wang et al.,
1999). Embryonic lethality was expected based on this antisense oligonucleotide
experiment in chick, however, viable and fertile mXinα knockout mice were observed.
This viability may result from functional compensation through the up-regulation of
mXinβ at both message and protein levels (Gustafson-Wagner et al., 2007). Consistent
with a possible compensatory role for mXinβ, mXinβ like mXinα (Sinn et al., 2002;
Wang et al., 1999), localizes to the intercalated disc of adult heart (Gustafson-Wagner et
al., 2007). Despite the expression of mXinβ, the adult mXinα-deficient mouse hearts are
hypertrophied and exhibit cardiomyopathy with conduction defects (Gustafson-Wagner et
al., 2007). This suggests that each of the mXin proteins may have a unique function in the
heart. However, the molecular mechanism behind mXinα functions remains to be
elucidated. The first step toward answering this question is to characterize the functional
domains on mXinα and its interacting partners.
Studies with the human homologs (hXinα also termed Cmya1 and hXinβ, also
termed Cmya3) of mXinα and mXinβ reveals that the Xin repeats bind actin filaments in
vitro, and that a minimum of 3 Xin repeats is required for the detectable binding
45
(Pacholsky et al., 2004). Therefore, Xin proteins should have multiple independent actinbinding sites, which could then cross-link actin filaments into a loosely packed
meshwork. This cross-linking activity has been demonstrated only with recombinant
protein containing 3~16 Xin repeats from hXinα (Cherepanova et al., 2006; Pacholsky et
al., 2004) but not with full-length hXinα protein. Additional studies with hXinα
demonstrated the ability of hXinα to directly bind to filamin c, Mena/VaSP, as well as to
colocalize at the intercalated discs, suggesting that Xin may play a role in remodeling of
the actin cytoskeleton (van der Ven et al., 2006). However, it is unclear why in the heart,
mXinα does not associate with actin thin filaments, but rather colocalizes with β-catenin
and N-cadherin at the intercalated disc. In the chicken heart, cXin is also associated with
the N-cadherin/β-catenin complex, as demonstrated by co-immunoprecipitation (co-IP)
experiments (Sinn et al., 2002). Thus, the molecular mechanisms underlying the role of
Xin in cardiac morphogenesis and myofibrillogenesis remain to be elucidated.
In this study, we gained further insight into the function of mXinα through the
identification of mXinα binding partners, including β-catenin and actin. Additionally, we
have mapped the β-catenin binding domain to aa residues #535-636 of mXinα which
overlaps with the actin binding domain. To investigate Xin’s involvement with the actin
cytoskeleton, we also studied the significance of Xin’s actin binding ability. Using
negative staining electron microscopy, we found that recombinant full-length mXinα
protein aggregates actin filaments into ordered actin bundles. Full-length mXinα interacts
more weakly with actin than a C-terminal deletion mutant lacking the β-catenin binding
domain and the C-terminus, but retaining most of the Xin repeats. The binding of mXinα
to actin filaments can be further enhanced by the presence of β-catenin. From these
results, we propose a model in which the β-catenin binding domain and the C-terminus of
mXinα prevent an interaction between full-length mXinα and actin (an auto-inhibited
state), until the β-catenin binding domain is occupied by β-catenin. The binding of
46
mXinα to β-catenin at the adherens junction of the intercalated disc would then enable
subsequent actin binding and bundling (an open state).
Materials and Methods
Yeast Two-Hybrid Assay and Library Screening
Protein-protein interactions between mXinα and either β-catenin or N-cadherin
were tested utilizing the Matchmaker Two-Hybrid System 3 (Clontech, Palo Alto, CA) in
yeast strain AH109. Full-length mXinα cDNA (Lin et al., 2005) was subcloned into the
SalI/SmaI sites of pEGFP-C2 (Clontech). The resulting plasmid pEGFP-mXinα was used
for constructing various deletions by PCR-based mutagenesis. The mXinα cDNA and
several deletion fragments were subcloned into the SalI/SmaI sites of the pGBKT7
vector. The full-length construct (pGBKT7-mXinα) encodes aa#1-1129 of mXinα. The
mXinαRΔ-1, -2 and -3 constructs (pGBKT7-mXinαRΔ-1, 2 and 3) represent mutants
with a deletion of the first half (aa#73-361), the second half (aa#362-746), and all (aa#73746), respectively, of the Xin repeats. The mXinαCΔ is a C-terminal deletion construct
(pGBKT7-mXinαCΔ) encoding aa#1-532. Another construct, pGBKT7-β-catBR,
encoding aa#533-746, was PCR amplified using the following primers:
5’AGTACCATCGATGTGGTACG3’, and 5’ AGCCCATGGGACAGTTTTC 3’. The
resulting product, after ClaI/NcoI digestion and fill in with the Klenow fragment, was
subcloned into the SmaI site of pGBKT7. This β-catenin binding region was further
divided into 4 fragments (fragment CA, AP, PN, and CP), each of them flanked with a
pair of ClaI, ApaI, PstI or NcoI sites. Primer pairs used to generate these fragments were:
5'-AGTACCATCGATGTGGTACG-3' and 5'-CAGAGAGATTGGGGCCCTTTCAT-3'
for the CA fragment, 5'-ATGTTTGGGCCCCAATCTCTG-3' and 5'CACCCGGCTGCAGTACCTTAC-3' for the AP fragment, 5'GTAAGGTACTGCAGCCGGGTG-3' and 5'-AGCCCATGGGACAGTTTC-3' for the
PN fragment and 5'-AGTACCATCGATGTGGTACG-3' and 5'-
47
CACCCGGCTGCAGTACCTTAC-3' for the CP fragment. The individual PCR
amplified fragment was then digested with the appropriate enzymes, filled in using the
Klenow fragment, and subcloned into the SmaI site of the pGBKT7 vector. The insert
sequences of all constructs were confirmed by DNA sequencing at the Roy J. Carver
Center for Comparative Genomics, Department of Biological Sciences, University of
Iowa.
To construct pGADT7-β-catenin, pGEX-KG-β-catenin (a generous gift from Dr.
Janne Balsamo, University of Iowa) was digested with BamHI, filled in, and subcloned
into the NdeI site of pGADT7. To construct pGADT7-N-cadherin, the N-cadherin
cytoplasmic domain was PCR amplified from the pSP72-N-cadherin (a gift from Dr.
Janne Balsamo) with the primers: 5’ggaattcATGAAGCGCCGTGATAAGG 3’ and
5’ccatcgatAATAAAAGCAATGCGATGTAAC 3’. This PCR fragment was digested and
subcloned into the EcoRI/ClaI sites of pGADT7. These prey constructs were separately
transformed into AH109, which had been previously transformed with either full length
mXinα or one of the deletion constructs. Direct interaction between the N-cadherin or βcatenin prey and the mXinα bait was determined using growth on selective media and a
β-galactosidase (β-gal) expression (X-gal assay) as described in the Two-Hybrid System
user manual (Clontech). The yeast two-hybrid assay was validated using p53 and the
large T antigen as a positive control. As a negative control, β-catenin was replaced with
the large T antigen.
In order to identify novel mXinα-interacting proteins, pGBKT7-mXinα was
further used as bait to screen a custom Matchmaker cDNA library prepared from 5 week
old rat hearts in the pGAD10 vector (Clontech Laboratories, Inc., Palo Alto, CA). After
recovery of positive prey plasmids and retransformation to confirm the interaction,
complete insert sequences were determined. Among 20 known and novel positive clones,
two independent clones, pL1.192 and pL2P6, encoding aa#1-258 and aa#1-215,
respectively, of rat cardiac α-actin as well as a clone, pL2P10, encoding aa#497-780 of
48
rat gelsolin were obtained and reported here as mXinα-interacting proteins. The other
obtained positive clones will not be addressed here. In other screenings using either a pretransformed mouse 17-day embryo Matchmaker cDNA library in the pACT2 prey vector,
or a custom adult mouse heart Matchmaker cDNA library constructed in the pGADT7RecAB prey vector, two independent clones, p4Q39 and p4Q79, each encoding fulllength mouse cardiac α-tropomyosin, and another clone, pL3Q8, encoding aa#2,5332,603 of mouse filamin b were obtained and reported here. In addition, full-length
cDNAs for p120 catenin (a generous gift from Dr. Janne Balsamo), talin (a generous gift
from Dr. Richard Hynes, MIT) and vinculin (a generous gift from Dr. Wolfgang
Goldman, University of Erlangen, Germany) were individually subcloned into pGADT7
plasmids and used as preys in yeast two-hybrid assay to test whether these proteins
interact with mXinα.
Constructions of Plasmids and Purification of Recombinant
Proteins
Expression plasmid pGEX-mXinα for GST-mXinα fusion protein was constructed
by ligating the EcoRI mXinα cDNA insert from pGBKT7-mXinα with EcoRI digested
pGEX4T-1 vector. Another expression plasmid, pET30-mXinα for His-mXinα fusion
protein was derived from the SalI/SmaI mXinα fragment (5.6kb) of pGEM3ZmXin3
subcloned into the XhoI(fill-in)/SalI sites of pET30a vector. pGEX-KG-β-catenin was
used for the production of GST-β-catenin fusion protein. Recombinant proteins were
expressed in E. coli BL21(DE3)pLysS cells and purified by Glutathione-sepharose 4B
column for GST-tagged proteins or His GraviTrap column for His-tagged proteins (GE
Healthcare Bio-Sciences Corp., Piscataway, NJ) according to the manufacturer’s
protocols.
49
Co-immunoprecipitation(Co-IP), Pull-down Assay and
Western Blot Analysis
Adult mouse hearts were homogenized in IB buffer: 20 mM phosphate buffer pH
7.5, 150 mM NaCl, 1% NP-40, 0.1% SDS and protease inhibitor cocktail (Roche,
Germany). The homogenate (150 µg total protein/immunoprecipitation) was cleared by
centrifugation at 12,000xg for 15 min and incubated with anti-β-catenin, anti-N-cadherin,
anti-p120 catenin, anti-plakoglobin, anti-filamin c, anti-vinculin or control mouse serum
for 2h, followed by protein G-sepharose beads (GE Healthcare) for 1h, at 4oC. The beads
were washed 3 times with IB buffer and once with PBS. The bound proteins were eluted
in SDS-PAGE sample buffer, fractionated by 7.5% SDS-PAGE and immunoblotted with
anti-mXinα U1013 antibody or anti-β-catenin as described previously for Western blot
analysis (Gustafson-Wagner et al., 2007).
For the pull-down assay, various amounts (5-60 nM) of recombinant His-mXinα
were mixed with 30 nM GST-β-catenin in binding buffer containing 20 mM HEPES
buffer pH 7.5, 100 mM KCl, 1 mM DTT (dithiothreitol), 0.1% Triton X-100 and 2.5 mM
PMSF (phenylmethanesulfonyl fluoride) for 2h at 4oC. The mixture was
immunoprecipitated with a monoclonal anti-β-catenin antibody. The immunoprecipitate
was further analyzed by Western blot with polyclonal anti-mXinα antibody as described
above.
Actin Binding Assay
Rabbit skeletal muscle actin (>99% pure) was purchased from Cytoskeleton, Inc.
(Denver, CO) and used in a slightly modified actin binding assay as described previously
(Novy et al., 1993). Briefly, Actin (9.3 µM) and various amounts of His-mXinα (0~7.64
µM) or GST-mXinα were mixed in 100 µl of 10 mM HEPES buffer pH 7.5, 100 mM
KCl, 0.05% Triton X-100, 0.1 mM DTT, 0.1 mM ATP and 1.5 mM MgCl2 in the absence
or presence (1.95 µM) of GST-β-catenin. The mixtures were incubated at room
50
temperature for 30 min and subjected to high speed centrifugation for 30 min in a
Beckman airfuge at 26 psi (100,000xg) to separate bound and unbound fractions. To test
the cross-linking or bundling activity of recombinant mXinα, the protein mixed with actin
was subjected to low speed centrifugation (10,000xg) for 15 min. Under this condition,
actin filaments remain in the supernatant except cross-linked or bundled filaments formed
by binding proteins. Aliquots of the supernatant and pellets were analyzed by 7.5% SDSPAGE and the protein bands were visualized by staining and quantified as described
(Novy et al., 1993).
Cell Culture, DNA Transfection and Fluorescence
Microscopy
The mouse skeletal muscle cell line, C2C12, was grown on glass coverslips in
DME low glucose medium plus 5% fetal bovine serum (FBS) and 15% defined
supplemented calf serum in a humidified incubator at 37oC with 5% CO2. Chinese
Hamster Ovarian (CHO) cells were grown on glass coverslips in DMEM plus 10% FBS.
Myoblasts or CHO cells were transfected with pEGFP-C2 (empty vector control),
pEGFP-mXinα, pEGFP-mXinαRΔ-1, -2, -3 or pEGFP-mXinαCΔ using Lipofectamine
PLUS reagent (Life Technologies, Rockville, MD) as previously described (Sinn et al.,
2002). After 24h, cells on coverslips were fixed in 3.7% formaldehyde and either
processed for immunofluorescence microscopy with a monoclonal anti-vinculin antibody
(Sigma) and a rhodamine-conjugated goat anti-mouse secondary antibody or directly
mounted onto glass slides and observed under a Zeiss epifluorescence photomicroscope
III. The fluorescence and phase-contrast images were collected with a Leica digital
camera and processed using Adobe Photoshop.
Electron Microscopy
Small aliquots (10µl) of actin and recombinant mXinα mixtures in actin binding
assay conditions were applied to carbon-coated Formvar grids and negatively stained
51
with 1.0% uranyl acetate. Samples were then observed under a JEOL 1230 transmission
electron microscope at an accelerating voltage of 100 kV (Central Microscopy Research
Facility, University of Iowa). The images were collected with Gatan CCD digital camera
attached to the electron microscope. The thickness of the actin bundles was measured
from images against a stained catalase resolution standard (Polysciences, Inc.,
Warrington, PA).
Results
mXinα is Associated with N-cadherin, β-catenin and p120
Catenin in the Adult Mouse Heart
The cXin gene is believed to play a vital role in cardiac morphogenesis (Wang et
al., 1999), possibly through an association with N-cadherin and β-catenin (Sinn et al.,
2002) and by intracellular signaling at the adherens junctions of the intercalated discs. In
the mouse heart, co-localization of mXinα with N-cadherin and β-catenin at the
intercalated discs has been demonstrated throughout embryogenesis and adulthood (Sinn
et al., 2002). Targeted deletion of mXinα in the mouse results in cardiac hypertrophy and
cardiomyopathy with abnormal intercalated disc ultrastructure (Gustafson-Wagner et al.,
2007). Although the underlying mechanism leading to cardiac defects remains unclear,
significantly reduced expression of N-cadherin, β-catenin and p120 catenin proteins is
observed in the mXinα-null mouse heart (Gustafson-Wagner et al., 2007). This suggests
an association of mXinα with these adherens junctional components.
To test this association, co-IP experiments were carried out using anti-N-cadherin,
anti-β-catenin and anti-p120 catenin antibodies with adult mouse heart extracts. Western
blot analysis with U1013 anti-mXinα antibody identified that both mXinα and the
alternatively spliced isoform mXinα-a were present in the total extract, as well as in the
anti-β-catenin immunoprecipitate, the anti-N-cadherin immunoprecipitate, and the antip120 catenin immunoprecipitate, but not in the control mouse serum immunoprecipitate
52
(Figure 2.1 A, left panel). As expected, anti-N-cadherin immunoprecipitate but not antiplakoglobin immunoprecipitate also contains β-catenin (Figure 2.1 A, right panel).
These data suggest that mXinα and mXinα-a are indeed components of the Ncadherin/β-catenin/p120 catenin complex and, therefore, supports our finding of
simultaneous down-regulation of these components in mXinα-null mouse heart
(Gustafson-Wagner et al., 2007). It should be noted that the mXinα-a isoform was
frequently over-represented in these immunoprecipitates relative to mXinα, suggesting
that these two isoforms may have slightly different mechanisms for association with the
immunocomplexes. As we have previously showed, the minor isoform, mXinα-a, is
encoded by an alternatively spliced mRNA with the inclusion of intron 2 from the mXinα
gene (Gustafson-Wagner et al., 2007). Therefore, the aa sequences of both mXinα-a and
mXinα are identical, except in the C-terminus, where the last two residues of mXinα are
replaced with an additional 683 aa residues. This extra C-terminal sequence was also
found in one (called Xin A) of the hXinα isoforms (van der Ven et al., 2006). A filamin
c-binding domain was previously mapped to the last 158 residues of this hXinα isoform,
Xin A (van der Ven et al., 2006). When the co-IPs were further performed with antifilamin c or anti-vinculin antibody, the resulting immunoprecipitates contained both
mXinα-a and mXinα (Figure 2.1 A left panel), suggesting that both mXinα isoforms are
associated with focal adhesion components, in addition to the N-cadherin/catenin
complex. The additional association between filamin c and mXinα-a in the extra Cterminal region may account for the observation that more mXinα-a is associated with
these immunoprecipitates, as mXinα lacks this region.
mXinα Directly Interacts with β-catenin
To examine whether a direct interaction between mXinα and β-catenin exists, coIPs with purified proteins (Figure 2.1B) and yeast two-hybrid assays (Figure 2.2) were
carried out. Increasing amounts of purified, recombinant His-mXinα were mixed with
53
GST-β-catenin in solution, followed by immunoprecipitation with monoclonal anti-βcatenin and Western blot analysis of the immunoprecipitate with U1013 anti-mXinα
polyclonal antibody. As can be seen in Figure 2.1 B, a 30 nM concentration of GST-βcatenin was able to co-precipitate increasing amounts (5 ~ 60 nM) of His-mXinα protein.
On the other hand, the immunoprecipitation in the absence of GST-β-catenin could not
bring down His-mXinα protein. These results clearly suggest a direct interaction between
mXinα and β-catenin. The amount of His-mXinα brought down by 30 nM GST-β-catenin
did not reach a plateau when 30 nM His-mXinα were added, further suggesting that
multiple mXinα-binding sites could be present in the β-catenin molecule, if mXinα
functions as a monomer.
A yeast two-hybrid assay was also used to determine whether mXinα directly
interacts with N-cadherin and β-catenin, and to map the domain of these potential
interactions, as we have shown that mXinα is part of the N-cadherin/β-catenin/p120
catenin complex (Figure 2.1 A). Full-length mXinα cDNA and several deletions were
constructed into the pGBKT7 vector and used as baits (Figure 2.2 A). β-catenin or the
cytoplasmic domain of N-cadherin in pGADT7 served as the “prey”. A direct proteinprotein interaction was not observed for mXinα and N-cadherin (data not shown).
However, as shown in Figure 2.2 B, mXinα, mXinαRΔ-1 and β-catBR, but no other
deletions, directly interact with β-catenin and give rise to positive X-gal stain. These
results demonstrate that the β-catenin binding region (β-catBR) on mXinα is localized
within the region from aa#533 to #746.
This region was shown to be both necessary and sufficient for the interaction with
β-catenin. Interestingly, this β-catenin binding region is located within the last 4 Xin
repeats of mXinα. As shown in Figure 2.2 C, this β-catenin binding region was further
divided into 4 fragments (CA, AP, PN, and CP) and used as baits in yeast two-hybrid
assay to map the minimal region for binding to β-catenin. Only CA and CP baits showed
positive interaction with β-catenin prey (Figure 2.2 C). Thus, the minimal β-catenin-
54
binding domain resides in the CA fragment, located within aa# 535 to #636. The aa
sequence alignment among Xin proteins from chick, mouse and human reveals high
sequence identity within this β-catenin-binding domain (Figure 2.2 D): 57.8% between
mXinα and hXinα, 53.9% between mXinα and mXinβ and 49.0% between mXinα and
cXin. The secondary structure of this β-catenin-binding domain predicted by ChouFasman method (Chou, 1990; Chou and Fasman, 1978) is composed of 39.2% and 35.3%
of β-sheet and α-helix amino acids, respectively. Each of them forms two stretches,
organizing into β-sheet (15aa) - α-helix (26aa) - β-sheet (21aa) - α-helix (14aa) structure.
mXinα Binds and Bundles Actin Filaments
It has been shown that recombinant Xin repeats from hXinα are capable of
binding and cross-linking actin filaments (Cherepanova et al., 2006; Pacholsky et al.,
2004). To test whether full-length mXinα also binds to actin filaments, recombinant
GST-tagged or His-tagged mXinα was used in an actin binding assay as described in
Experimental Procedures. After high-speed centrifugation (100,000xg for 30 min), both
GST-mXinα and His-mXinα are co-sedimented with actin filaments into the pellet
fraction (P in Figure 2.3 A tubes# 1-6). Under the same conditions, GST-mXinα or HismXinα alone remains in the supernatant (S in Figure 2.3 A tubes# 7-8). GST-mXinα cosedimented with actin filaments appears to be more efficient than His-mXinα. Although
the exact mechanism for the higher efficiency remains unknown, the fact that GST alone
is able to form dimers in solution (Ji et al., 1992) is one possible explanation. The binding
of His-mXinα to actin filaments appears to be saturable (Figure 2.3 B). The estimated
molar ratio of actin to bound mXinα is 9.1:1 at saturation. Purified actin filaments in
solution could not be pelleted by low-speed centrifugation (10,000xg for 15 min) unless
the filaments become aggregates by actin cross-linking or bundling proteins. Figure 2.4
shows the results of such low-speed co-sedimentation assay. Increasing amounts of
recombinant GST-mXinα cause increasing actin filament aggregation (Figure 2.4, lanes
55
2~6P). Under these conditions, GST-mXinα alone remained in the supernatant (Figure
2.4, lane 7S), actin filaments alone could not be sedimented (lane 1P), and recombinant
GST by itself (lane 8P) or bovine serum albumin (BSA) (lane 9P) also could not
aggregate actin filaments into the pellet (Figure 2.4). Similarly, we found that His-mXinα
was able to aggregate actin filaments, as determined by low-speed co-sedimentation
assay (Figure 2.5). These results suggest that full-length mXinα is capable of either crosslinking or bundling the actin filaments.
To distinguish between these two activities, negatively stained actin filaments in
the absence and presence of recombinant mXinα were examined under the electron
microscope. Actin filaments alone are thin and long with average diameter of 6~8 nm
(Figure 2.6 A). At a ratio of actin to recombinant His-mXinα of 5:1, actin filaments were
aggregated into side-by-side bundles (Figure 2.6 B). Similar bundling activity was also
observed with GST-mXinα (Figure 2.6 C). Furthermore, the size of the bundles which
form increase with the concentration of His-mXinα. At a 10:1 molar ratio of actin to HismXinα (black bars in Figure 2.6 D), the average bundle sizes formed after 13 and 48 hrs
of incubation are significantly smaller than that formed at the ratio of 5:1 (shaded bars in
Figure 2.6 D, p<0.05), suggesting that the bundling reaction is mXinα concentration
dependent. Although there was no transverse band observed in these actin bundles,
individual actin filaments appeared to be decorated by mXinα with a periodicity of
36.0±0.4 nm (n = 129) (Figure 2.6 E) at higher actin to mXinα ratios of 1:2. The nature
and significance of this periodicity remain to be determined.
The β-catenin-Binding Domain and the C-terminal Half of
mXinα Prevent Ectopically Expressed mXinα from
Localizing to Stress Fibers within C2C12 Myoblasts
Recombinant mXinα and its Xin repeat region bind and aggregate actin filaments
in vitro; however, in the adult mouse heart, the mXinα protein does not associate with
56
actin thin filaments, instead, mXinα preferentially localizes to the intercalated discs.
Undifferentiated C2C12 myoblasts do not express mXinα (Lin et al., 2001); however,
upon differentiation, mXinα expression in myotubes is localized to a few stress fibers and
near the periphery of the cell (Sinn et al., 2002). The protein domain responsible for its
localization was investigated in the C2C12 cells by transient transfections with plasmids
expressing GFP-mXinα or its various deletion constructs. Control C2C12 myoblasts
transfected with pEGFP-C2 vector alone showed diffusely distributed GFP at the
perinuclear region and in the nucleus (Figure 2.7 A). In contrast, myoblasts transfected
with the pEGFP-mXinα exhibited some GFP-mXinα fusion protein localized to stress
fibers (Figure 2.7 B), and cells transfected with pEGFP-mXinαRΔ-3, which lacks all 15
Xin repeats and the β-catenin-binding domain, showed a diffuse distribution of GFPmXinαRΔ-3 with very little stress fiber localization (Figure 2.7 C). When the C-terminal
deletion construct was used, pEGFP-mXinαCΔ, which lacks the β-catenin-binding
domain as well as the C-terminal proline-rich region, transfected cells demonstrated
increased stress fiber localization and peripheral localization of GFP-mXinαCΔ (Figure
2.7D). Nearly 100% of pEGFP-mXinαCΔ transfected myoblasts had clearly visible stress
fiber and peripheral staining, whereas <50% of myoblasts transfected with other plasmids
exhibited such localization. All transfected proteins associated with stress fibers
colocalized with either vinculin or phalloidin within myoblasts (data not shown).
Interestingly, cells transfected with pEGFP-mXinαRΔ-3, lacking all 15 Xin repeats and
β-catenin-binding domain, appeared to have inhibited cell spreading ability, resulting in a
smaller apparent cell size. The differences in cell size and shape were further
characterized in transiently transfected CHO cells using the 2D dynamic image analysis
software (2D-DIAS) program (Li et al., 2004; Soll, 1995; Soll and Voss, 1998).
Transfection of the full-length mXinα (pEGFP-mXinα) did not appear to affect cell size
and shape compared to the control pEGFP-C2 vector transfection, as measured through
mean cell area (µm2), perimeter (µm) or roundness (%) (Table 2.1). However, pEGFP-
57
mXinαRΔ-3 significantly reduced cell area and roundness relative to the full-length or
control vectors (Table 2.1). These results suggest that the β-catenin-binding domain
together with the C-terminus of mXinα may inhibit stress fiber localization of expressed
GFP-mXinα. The Xin repeats appeared to be important for cell shape and size (spread
area) determination.
Function of the Xin Repeats in Stress Fiber Localization
In order to determine the function of the Xin repeats in mXinα, CHO cells
expressing GFP-mXinα or to various deletion constructs were counterstained with
monoclonal anti-vinculin antibody and rhodamine-conjugated secondary antibody and
analyzed by fluorescence microscopy. A representative GFP image and a merged image
with vinculin localization from each transfected cell line is shown in Figure 2.8. The
vinculin staining at the focal adhesion site was used to confirm the stress fiber
localization of GFP fusion proteins. It is clear that an increasing association of GFP
fusion protein with stress fibers is seen in the following order: GFP-mXinαRΔ-3 (Figure
2.8 A,B) < GFP-mXinαRΔ-1 (Figure 2.8 C,D) < GFP-mXinαRΔ-2 (Figure 2.8 E,F) <
GFP-mXinα (Figure 2.8 G,H) < GFP-mXinαCΔ (Figure 2.8 I,J). These results are
consistent with observations from transfected C2C12 myoblasts (Figure 2.7).
For quantification, randomly selected cells from each transfected cell line were
scored for the frequency of GFP-signal associated with detectable stress fibers, and then
grouped into 4 categories: group I through IV (cells with 0, 3-9, 10-20 and >20 stress
fibers, respectively). As shown in Table 2.2, deletion of the last 7 Xin repeats in
mXinαRΔ-1 leads to a weaker stress-fiber association (a significant increase in group II)
compared to the wild-type mXinα construct with the 15 Xin repeats (p<0.01, chi square
test). However, deletion of the first 8 Xin repeats (mXinαRΔ-2) results in an even weaker
association with stress fibers (a significant decrease in group II, p<0.025, and a
significant increase in group III, p<0.0001). Expression of mXinαRΔ-3, which
58
completely lacks the Xin repeats as well as the overlapping β-catenin-binding domain,
totally abolishes the stress-fiber association (almost all cells are categorized into group I).
Thus, the extent of stress fiber association is roughly proportional to the number of the
Xin repeats present in these expressed proteins. However, the mXinαCΔ construct, which
contains 10 Xin repeats but lacks the β-catenin-binding domain and the C-terminus of the
mXinα protein, exhibits an even stronger stress fiber association than the wild-type
mXinα with 15 Xin repeats (increased group IV cells, p<0.025). This result again
suggests that the β-catenin-binding domain and the C-terminus of mXinα may play an
inhibitory effect on actin association.
It is known that a minimum of 3 Xin repeats is required for actin binding
(Pacholsky et al., 2004). Therefore, a comparison between force-expressed mXinαRΔ-1
with 7 Xin repeats and mXinαRΔ-2 with 8 repeats, or between mXinαRΔ-2 with 8 Xin
repeats and mXinαCΔ with 10 repeats, would not be expected to have a major effect on
their stress-fiber association. The major difference between mXinαRΔ-1 and mXinαRΔ-2
or between mXinαRΔ-2 and mXinαCΔ is in the presence or absence of the β-cateninbinding domain and the C-terminus, respectively. The observed, significant differences in
stress fiber association for these two comparisons (p valueb and p valuec in Table 2.2) is
consistent with the inhibitory role of the β-catenin-binding domain and the C-terminus in
the ability of mXinα to bind actin.
In a separate yeast two-hybrid assay to identify novel mXinα interaction partners,
a rat heart cDNA yeast two-hybrid library was screened using mXinα as bait. From this
screen, we obtained two independent clones, pL1.192 and pL2P6, encoding aa#1-258 and
aa#1-215, respectively, of cardiac α-actin. Interestingly, the interaction between actin and
mXinαCΔ appeared to be stronger, as observed with more β-galactosidase activity
(stronger blue), than that between actin and mXinα in yeast cells grown on nutritionally
selective plates containing X-gal (Figure 2.9 A). Using liquid culture assay with ONPG
as substrate for β-galactosidase to quantify the amounts of reporter gene expression after
59
bait and prey interaction in yeasts (Clontech Two-Hybrid system user manual), we found
about 3.6-fold higher β-galactosidase expression in cells with mXinαCΔ and actin
relative to cells with mXinα and actin. These results are in good agreement with the idea
that the β-catenin-binding domain and the C-terminus of mXinα prevent mXinα’s binding
to actin filaments.
In addition to two cardiac α-actin clones obtained from yeast two-hybrid library
screening, we have also identified several positive clones, encoding fragments of known
actin-binding proteins, including filamin b (aa#2,533-2,603), muscle α-tropomyosin (fulllength aa#1-294) and gelsolin (aa#497-780). Re-transformation of these prey plasmids
with the mXinα bait in yeast confirmed the interactions, as all showed blue color in a βgalactosidase filter assay (Figure 2.9 B). Under the same conditions, full-length p120
catenin and full-length vinculin preys exhibited a positive interaction with mXinα
whereas full-length talin prey, employed as a negative control, did not interact with
mXinα (Figure 2.9 B).
The Presence of β-catenin Enhances mXinα Binding to
Actin Filaments In Vitro
We have shown that the β-catenin binding domain overlaps with the 12th and 13th
Xin repeats, and that the β-catenin binding domain and the C-terminus of mXinα together
reduce mXinα binding to actin stress fibers. These results led us to ask whether the
binding of β-catenin to mXinα would release the inhibition and then enhance the actin
binding. To address this question, we performed an actin binding co-sedimentation assay
and a quantitative measurement of sizes of actin bundles formed by His-mXinα in the
absence and presence of GST-β-catenin. As expected, in the absence of GST-β-catenin,
the amounts of His-mXinα co-sedimented with actin filaments increased as increasing
amounts of His-mXinα were used in the co-sedimentation assay (Figure 2.10). In the
60
presence of β-catenin, the amounts of His-mXinα co-pelleted with actin appeared to
increase about 2 fold (Figure 2.10).
The effect of β-catenin on mXinα actin bundling activity was further analyzed by
negative staining electron microscopy. In this experiment, His-mXinα alone or an equal
molar mixture of His-mXinα and GST-β-catenin were pre-incubated on ice for 2 hrs and
then mixed with actin filaments. At different time points (1.5, 8 and 30 min.), aliquots
from the mixture were applied onto formvar grids and processed for negative staining.
Samples were observed under JEOL-1230 electron microscope and 50 micrographic
pictures were randomly taken from each sample. The width of all the bundles from the
micrographs was quantified using the ImageJ program. Substitution of GST-β-catenin in
the mixture by GST was used as a negative control. With an increase in the incubation
time, actin bundles formed by mXinα alone or a combination of mXinα and β-catenin
increased in both numbers and sizes (Figure 2.11). GST did not have significant effects
on mXinα’s actin bundling activity (data not shown). However, it is clear that in the
presence of β-catenin, many more bundles were formed at given time point and also
bigger bundles appeared more frequently (Figure 2.11). Thus, these results together show
that β-catenin enhances actin bundling by mXinα, and is consistent with a model in which
β-catenin binding to mXinα releases the inhibition of actin binding imposed by the βcatenin-binding domain and the C-terminus, and then facilitates the mXinα’s actin
binding/bundling activity.
Discussion
Model for How mXinα Functions at the Adherens Junction
of the Heart
In the present study, we have shown that β-catenin is capable of directly
interacting with aa#535-636 of mXinα. Through this interaction, mXinα can subsequently
enhance its ability to bind and to bundle actin filaments. Furthermore, through this
61
interaction, mXinα can then associate with the N-cadherin/β-catenin/p120 catenin
adhesion complex. That mXinα plays a pivotal role in vivo, is evident from the cardiac
defects observed in the mXinα-null mice (Gustafson-Wagner et al., 2007). The finding
that the β-catenin-binding domain on mXinα overlaps with the actin binding Xin repeats
provides a plausible explanation for why mXinα is not associated with the actin filaments
of sarcomeres, even though it contains 15 Xin repeats, which constitute multiple actin
binding sites. Rather, mXinα is preferentially localized to the intercalated disc of the
heart.
The results presented here imply that newly synthesized mXinα may be present in
an auto-inhibited state, as far as actin binding is concerned, until the β-catenin-binding
domain is occupied by the β-catenin. The binding of mXinα to β-catenin at the adherens
junction may change the conformation of mXinα into an open state which may enable
subsequent actin binding and bundling (Figure 2.12). In this regard, mXinα could be
considered an integral component of adherens junctions at the intercalated disc, where it
links the N-cadherin-mediated adhesion complex to the actin cytoskeleton. In the present
study, we have also provided strong evidence to support the proposed two state model for
mXinα interaction with actin. First, a deletion of the β-catenin-binding domain and the Cterminus found in mXinαCΔ mutant allows the Xin repeats to more readily bind to actin
and relieves the inhibition caused by the C-terminal deletion fragment. As a result,
mXinαCΔ strongly enhances its ability to interact with actin in yeast and to associate with
actin stress fibers in both transfected C2C12 and CHO cells. Second, in the presence of βcatenin, both actin binding and bundling activities are significantly increased.
Adherens junctions at the intercalated disc function to connect the mature
myocytes and to provide the attachment sites at the membrane for myofibrils (Ong et al.,
1998). This function is dependent upon the assembly of the cadherin-catenin complex
(Hertig et al., 1996). The highly conserved cytoplasmic domain of N-cadherin binds to βcatenin and/or plakoglobin (γ-catenin). p120-catenin, a distant relative of β-catenin, binds
62
to the juxtamembrane region of cadherin and regulates cadherin turnover (Reynolds and
Roczniak-Ferguson, 2004). In a classic view, an actin bundling protein, α-catenin, then
binds β-catenin to organize the adhesion complex that links to the actin cytoskeleton,
either via a direct association with actin filaments or through an indirect association with
actin binding proteins, including vinculin and α-actinin (Pokutta and Weis, 2002). This
stable linkage role for α-catenin has been recently proven to not exist in epithelial cells;
instead, compelling evidence suggests that α-catenin is a molecular switch that binds Ecadherin-β-catenin and regulates actin dynamics at the adherens junctions of epithelial
cells (Drees et al., 2005; Gates and Peifer, 2005; Yamada et al., 2005). The component
which then in fact makes this connection in epithelial cells remains unclear.
In this study, we have shown that the mXinα protein is a potent actin bundling
protein, which can additionally interact with β-catenin. Furthermore, β-catenin effectively
enhances the binding of mXinα to actin filaments. mXinα localizes to the adherens
junction in the intercalated disc of the heart (Sinn et al., 2002). Thus, it is possible that
mXinα provides a link between N-cadherin and actin cytoskeleton in cardiomyocytes.
Supporting a linkage role for mXinα, mXinα-null mouse hearts begin with abnormal
intercalated disc ultrastructure as early as 3 months of age and exhibit cardiac
hypertrophy and cardiomyopathy (Gustafson-Wagner et al., 2007). This structural
alteration is accompanied by a disorganization of myofibrils at the intercalated disc and
by a significant decrease in the expression of N-cadherin, β-catenin and p120-catenin
(Gustafson-Wagner et al., 2007), suggesting that hypertrophy may be due to impaired
organization of the intercalated disc and instability of cell-cell adhesion. Although the
lack of mXinα protein in the intercalated discs is the most straightforward explanation for
the observed cardiac defects, there are other possibilities, such as the up-regulation of
mXinβ in mXinα-null heart, for partially contributing to the observed phenotype. We
have shown here that over-expression of mutant mXinαRΔ-3 protein but not the wildtype mXinα protein in CHO cells altered cell size and shape. Therefore, it is unlikely that
63
the up-regulation of wild-type mXinβ in the mXinα-null heart would be the major factor
causing the observed cardiac defects.
mXinα Contains a Novel β-catenin-Binding Domain
The amino acid sequence from #613 to #685 of mXinα, which overlaps slightly
with the identified β-catenin-binding domain (aa#535-636), shows a 30% sequence
identity (39% similarity) to aa#2237-2305 of adenomatous polyposis coli (APC), a
known β-catenin binding protein involved in canonical Wnt signaling (Nelson and Nusse,
2004). Despite this similarity, APC does not use this region to bind β-catenin. Instead, the
β-catenin-binding domain on APC has been mapped to aa#1014-1210, containing 3 of
15-aa repeats (Rubinfeld et al., 1993; Shih et al., 2000; Su et al., 1993). Moreover, the βcatenin-binding domain of mXinα is different from the β-catenin-binding domains found
in axin (Behrens et al., 1998; Ikeda et al., 1998; Xing et al., 2003), α-catenin (Pokutta and
Weis, 2000), N-cadherin (Sadot et al., 1998), and Tcf (Behrens et al., 1996; Molenaar et
al., 1996). Therefore, mXinα possesses a novel β-catenin-binding domain. As shown in
Figure 2.2 D, this β-catenin-binding domain on mXinα is highly conserved among all Xin
proteins (49~57.8% identity). In the chicken heart, cXin has been shown to localize to the
intercalated disc (Lin et al., 2001) and to associate with the N-cadherin/β-catenin
complex (Sinn et al., 2002). We have also shown that the mXinβ messages and proteins
localize to the intercalated disc of the mouse heart (Gustafson-Wagner et al., 2007).
Moreover, up-regulation of mXinβ at the message and protein levels associated with the
targeted deletion of mXinα is consistent with a possible functional compensation between
mXinα and mXinβ in the heart (Gustafson-Wagner et al., 2007). Thus, it is possible that
the β-catenin-binding domains on cXin and mXinβ exist and are also functional.
mXinα Bundles Actin Filaments
A previous study with recombinant His-Xin repeats from hXinα has demonstrated
that 3 Xin repeats are necessary and sufficient to bind actin filaments, and that the Xin
64
repeats are capable of aggregating actin filaments into loosely packed meshwork
(Cherepanova et al., 2006; Pacholsky et al., 2004). This suggests that the Xin repeats
cross-link actin filaments. At saturation conditions, actin to Xin-repeat fragment molar
ratio is 4:1 for 16 repeats, 2:1 for 6 repeats or 1:1 for 3 repeats (Pacholsky et al., 2004).
However, using recombinant full-length mXinα in this study, we clearly demonstrate that
mXinα aggregates actin filaments into bundles, which can be sedimented by low speed
centrifugation (10,000xg for 15 min). This bundling activity is not attributable to the
fusion tags on mXinα. Both GST- and His-mXinα show similar bundling activity and
GST by itself does not have any actin binding/bundling activity. At saturation, one
molecule of His-mXinα binds to 9 molecules of actin monomers. The bundling reaction
of His-mXinα appears to be concentration- and time-dependent, and unlike another actin
cross-linking protein, filamin (Stossel et al., 2001), His-mXinα does not crosslink actin
filaments into a loosely packed network. His-mXinα fails to cross-link actin filaments in
all the concentration ranges we tested (the molar ratios of mXinα to actin tested ranging
from 2:1 to 1:200) (Figure 2.13). The differences in observed actin binding properties
(stoichiometry and cross-linking versus bundling) between the Xin repeats alone and fulllength mXinα suggest that a small portion (aa#1-72) of the N-terminal fragment upstream
of the Xin repeats and/or the C-terminal fragment (aa#747-1129) downstream of the Xin
repeats are also important in terms of organizing the actin cytoskeleton. Neither actin
polymerization activity nor G-actin binding activity is associated with recombinant Xin
repeats (Pacholsky et al., 2004). However, at the present time we do not know whether or
not full-length mXinα has these activities.
In yeast two-hybrid cDNA library screenings, we obtained two clones encoding
cardiac α-tropomyosin, one clone encoding gelsolin, and one clone encoding filamin b, in
addition to two clones encoding different fragments of cardiac α-actin. These proteins are
known to be actin binding proteins, either functioning to stabilize or regulate actin
dynamics and organization in cells (Pollard et al., 2000). Therefore, it is possible that
65
full-length mXinα functions through these interacting and actin binding proteins to
regulate its bundling activity. In the present study, we have shown that mXinα interacts
with filamin b, which is more broadly expressed in many tissues and cells, including
heart and C2C12 cells (van der Flier et al., 2002), than striated muscle-enriched filamin c
(Stossel et al., 2001). Since the mXinα-a isoform has exactly the same sequence as
mXinα, with the exception of a substitution of 2 residues at the C-terminus with 683
residues (Gustafson-Wagner et al., 2007), we expect that mXinα-a also interacts with
filamin b. The mXinα/mXinα-a binding site on filamin b appears to be located within
aa#2,533-2,603 at the end of the filamin b protein, a region that has high amino acid
sequence identity (70%) to filamin c. Previously, using muscle-specific Ig domain 20
from filamin c as a bait in yeast two-hybrid screens, van der Ven et al. identified a
binding partner containing the last 158 amino acid residues from the hXinα-a (human
homolog of mXinα-a) (van der Ven et al., 2006). However, this sequence is not present
the in mXinα isoform. Therefore, mXinα can only interact with filamin b, whereas
mXinα-a can use the same site to bind both filamin b and filamin c, as well as use the
extra site to bind muscle-specific filamin c.
The molecular mechanism of actin bundling by full-length mXinα remains to be
determined. Chemical cross-linking assays have revealed that recombinant Xin repeats by
themselves do not form polymers to cross link actin filaments (Pacholsky et al., 2004).
On the other hand, electron microscopy and iterative helical real space reconstruction to
visualize complexes of actin filaments with recombinant Xin repeats, showed that the Xin
repeats can bind to actin filaments in two distinct modes. In the side mode, residues from
subdomains three and four of one actin protomer and residues from subdomains one and
two of the adjacent actin protomer were the Xin-contact sites and in the front mode,
residues 22-27 and 340-345 of subdomain one provide the Xin-binding sites. Thus, each
actin molecule contains multiple binding sites for the Xin repeats (Cherepanova et al.,
2006), which together with multiple Xin repeats in mXinα would allow mXinα to bundle
66
actin filaments, like nebulin (Gonsior et al., 1998; Lukoyanova et al., 2002) and nebulette
(Cherepanova et al., 2006). The data presented here provides insight into the role of
mXinα in organizing the actin cytoskeleton and in linking between the adherens junction
and the actin cytoskeleton in cardiomyocytes. Future studies investigating these and other
mXinα interacting and actin-binding proteins may shed additional light on the precise
molecular mechanisms by which mXinα functions in the heart.
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Figure 2.1. Co-immunoprecipitation (Co-IP) of mXinα and β-catenin from adult mouse
heart and from purified recombinant proteins. (A) Total extract prepared from adult
mouse hearts was immunoprecipitated with mouse control serum, monoclonal anti-βcatenin, anti-N-cadherin, anti-p120 catenin, anti-filamin c, anti-vinculin, or antiplakoglobin. Western blots on immunoprecipitates were probed with the indicated
antibody. Both mXinα and its isoform, mXinα-a are detected in the anti-β-catenin, antiN-cadherin, anti-p120-catenin, anti-filamin c, and anti-vinculin immunoprecipitates, but
not in the control serum immunoprecipitate. β-catenin is detected in the anti-N-cadherin
immunoprecipitate but not the anti-plakoglobin immunoprecipitate. (B) Increasing
amounts of purified recombinant His-mXinα were mixed with GST-β-catenin in binding
buffer and subjected to immunoprecipitation (IP) by anti-β-catenin antibody. Western
blots (Blot) on the immunoprecipitate were probed with polyclonal anti-mXinα U1013
antibody to detect co-immunoprecipitated mXinα or anti-β-catenin antibody to
demonstrate equal amounts of β-catenin in the immunoprecipitate. An increasing amount
of mXinα directly binds to β-catenin. When an mXinα (60 µM) to β-catenin molar ratio
of 2:1 was used, the co-pelleted mXinα still increased. On the other hand, the mXinα
alone (control) at this mXinα concentration could not be co-pelleted. This result suggests
that a molecule of β-catenin can bind at least 2 molecules of mXinα.
68
69
Figure 2.2. Determination of the β-catenin-binding domain on mXinα. (A) Schematic
representation of mXinα and various deletion constructs used in the yeast two-hybrid
assay. The mXinα protein contains a putative DNA-binding domain (DBD), a putative
nuclear localization signal (NLS), a region with 15 Xin repeats, and a proline-rich region.
The deletion constructs include mXinαRΔ-1, mXinαRΔ-2, mXinαRΔ-3 (which represent
deletions of the first 8 repeats, the second half, and all 15 Xin repeats, respectively), as
well as mXinαCΔ (a C-terminal deletion containing only the first 10 repeats), and βcatBR (β-catenin-binding region). The relative strength of interaction with β-catenin, as
determined by an X-gal assay are indicated by the numbers of + symbols, and the –
symbol represents weak or no interaction. (B) Results of β-galactosidase filter assay for
the interaction between mXinα and β-catenin. After co-transformation of bait and prey
into yeast AH109 cells, colonies grown on selective media were transferred to a
membrane for the X-gal assay. The constructs which lack aa#533-746, including
mXinαRΔ-2 (c), mXinαRΔ-3 (d), mXinαCΔ (e), show low β-gal activity, indicating lack
of interaction. In contrast, the mXinα (a), mXinαRΔ-1 (b), and β-catBR (f) constructs,
which retain this region, demonstrate a high level of β-gal activity, suggesting a strong
interaction. The region represented by amino acids #533-746 is both necessary and
sufficient for the binding of mXinα to β-catenin. (C) The β-catenin binding domain
resides within the CA fragment of β-catBR. The β-catBR region of aa#533-746 was
further divided into 4 fragments (CA, AP, PN and CP). Each fragment was subcloned and
used as bait in yeast 2 hybrid assay. Only CA and CP containing a common region of aa
#535-636 showed strong interaction with β-catenin in the X-gal filter assay. Therefore,
the β-catenin binding domain locates within aa#535-636. (D) Comparison of the βcatenin binding domain of Xin proteins from mouse, human and chick. The β-catenin
binding domain (aa#535-636) defined on mXinα was used to align all predicted Xin
proteins from mouse (mXinβ: AY775570-775572), human (hXinα/Cmya1: AJ626899,
hXinβ/Cmya3: XM_496606) and chick (cXin: AF051944) using ClustalW program
(DNASTAR, Inc., Madison, WI). Dashes indicate gaps introduced for optimal alignment.
Yellow, green and red colors signify 5/5, 4/5 and 3/5 identity, respectively. This βcatenin-binding domain represents a highly conserved amino acid sequence among these
Xin proteins, and the sequence identity between mXinα and cXin, mXinβ or hXinα is
49.02%, 53.92% or 57.84%, respectively.
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Figure 2.3. Actin binding of purified recombinant His-mXinα and GST-mXinα. (A) SDSPAGE analysis of GST-mXinα and His-mXinα. An actin binding assay was performed
based on the co-sedimentation method at 100,000xg centrifugation. Under this condition,
both GST-mXinα (tube #7) and His-mXinα (tube #8) alone remained in the supernatant
(S). In the presence of muscle actin filaments (9.3 µM), an increasing amount of GSTmXinα (tube #1-3) and His-mXinα (tube #4-6) was co-sedimented in the pellet (P).
Numerous minor fragments recognized by anti-mXinα antibody (data not shown)
represent degraded products during purification. Particularly, a 100kDa band from GSTmXinα and a 80kDa band from His-mXinα are also co-pelleted with actin filaments. The
apparent difference in binding affinity between GST-mXinα and His-mXinα may be due
to the ability of GST by itself to form dimers (Ji et al., 1992) (B) Actin binding curve of
His-mXinα. The binding of His-mXinα to actin was determined by quantifying stained
gels of both supernatants and pellets. The amounts of bound His-mXinα per mole of actin
were calculated and plotted against total concentrations of His-mXinα using the
SigmaPlot 9.0 computer program.
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Figure 2.4. SDS-PAGE analysis of actin aggregates by GST-mXinα. An actin binding
assay was performed with increasing amounts of GST-mXinα as described under the
Figure 2.3 legend, except that low-speed centrifugation (10,000xg, for 15 min) was used.
Top gel represents pellet fraction (P), while bottom gel represents supernatant fraction
(S). Under this centrifugation condition, a trace amount of actin was pelleted from the
tubes containing actin alone (lane 1P), actin plus 2 µg GST-mXinα (0.13 µM, lane 2P),
actin plus GST (lane 8P) or actin plus bovine serum albumin (BSA) (lane 9P). However,
GST-mXinα at 5 µg (0.34 µM, lane 3P) started to bring down a detectable amount of
actin aggregates. The more GST-mXinα (10µg in lane 4P and 20 µg in lane 5P, 0.67 µM
and 1.34 µM respectively) added, the more actin aggregates were present. GST-mXinα at
30 µg (2.01 µM, lane 6P) did not bring down more actin aggregates, suggesting a
saturation point was reached. Under this assay condition, 30 µg GST-mXinα alone
remained in the supernatant (lane 7S).
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Figure 2.5. SDS-PAGE analysis of low speed actin co-sedimentation with His-mXinα.
The actin binding assay was performed with increasing amounts of His-mXinα as
described under Figure 2.4 legend. No detectable actin was pelleted from the tubes
containing actin alone (lane 1P), actin plus 2.5 µg His-mXinα (0.20 µM, lane 2P) or actin
plus bovine serum albumin (BSA) (lane 6P). His-mXinα at 10 µg (0.80 µM, lane 3P), 25
µg (2.0 µM, 4P) and 50 µg (4.0 µM, 5P) aggregated actin and can be co-pelleted down
with actin by low speed centrifugation. Consistent with the result described in Figure 2.4,
the reaction between His-mXinα and actin reached saturation when 25 µg His-mXinα
was mixed with actin (4P and 5P).
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Figure 2.6. Characterization of actin bundles formed by His-mXinα and GST-mXinα.
Electron microscopic images of actin alone (A), actin bundles formed by His-mXinα (B)
and actin bundles formed by GST-mXinα (C). From A to C, actin alone (2.4 μM) or actin
mixed with recombinant mXinα (0.48 μM) was applied onto a grid, negatively stained by
1% uranyl acetate and then observed under an electron microscope. Bar = 100 nm. (D)
The sizes of actin bundles formed by two different concentrations of His-mXinα were
measured from randomly selected micrographs and compared. The average bundle size
(width in nm) was affected by the concentration of mXinα in the mixture, even though
actin concentration was kept the same (2.4 μM). black bar, 0.24 μM His-mXinα; shaded
bar, 0.48μM His-mXinα; * indicates p<0.05 by rank sum test, the error bar represents
standard error. (E) Actin filaments that had not been included into the bundles were
decorated by His-mXinα, forming 36 nm periodicity marked by arrows. Bar = 100 nm.
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Figure 2.7. Immunofluorescence microscopy of transiently transfected C2C12 myoblasts.
Myoblasts maintained under growth conditions were transiently transfected with pEGFPC2 vector alone (A), pEGFP-mXinα (B), pEGFP-mXinαRΔ-3 (C), or pEGFP-mXinαCΔ
(D). Twenty-four hours post-transfection, cells were fixed and processed for fluorescence
microscopy to view GFP-tag protein expression. Expressed GFP-mXinα showed diffuse
staining, peripheral and stress fiber localizations (B), whereas most of GFP-mXinαRΔ-3
(lacking all 15 Xin repeats) were diffusely distributed within the cells with a very few
stress fiber localization (C). On the other hand, cells transfected with pEGFP-mXinαCΔ
(having 10 Xin repeats but lacking β-catenin-binding domain and the C-terminus)
showed much more GFP-signals associated with stress fibers and cell periphery than
wild-type GFP-mXinα (D), suggesting a possible inhibition of stress fiber localization by
β-catenin-binding domain and the C-terminus. The GFP-signal in the cells transfected
with empty vector was observed mainly in the nucleus and perinuclear regions (A). All
images are of the same magnification, Bar = 50µm.
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Figure 2.8. Immunofluorescence microscopy of CHO cells transfected with GFP-fulllength mXinα or various GFP-mXinα deletion constructs (mXinαRΔ-1, mXinαRΔ-2,
mXinαRΔ-3, mXinαCΔ). Transfected cells were processed for immunofluorescence
microscopy by counter-staining with anti-vinculin primary antibody and rhodamineconjugated secondary antibody. Because vinculin is known to localize to the focal
adhesion, anti-vinculin stains and GFP signals highlight the presence of the stress fibers
(B,D,F,H, and J). Cells expressing either the GFP-mXinα (G,H) or the GFP-mXinαCΔ
(I,J) contained many observable stress fibers. In contrast, while cells expressing the GFPmXinαRΔ-1 (C,D) or the GFP-mXinαRΔ-2 (E,F) exhibited a moderate decrease in the
number of stress fibers observed. Most notably, cells expressing the GFP-mXinαRΔ-3
(A,B) were much smaller in size and exhibited a significant reduction in the number of
observable stress fibers. These data are consistent with the quantitative data presented in
Table 2. Bar = 25 µm.
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Figure 2.9. Yeast two-hybrid assay to demonstrate the interaction of mXinα with mXinαinteracting proteins. Using mXinα as bait to screen yeast two-hybrid cDNA library,
cardiac α-actin and several actin binding proteins were identified as mXinα-interacting
proteins. (A) The interaction between C-terminal deletion mutant mXinαCΔ and actin is
stronger than that between full-length mXinα and actin. Yeast cells re-transformed with
either mXinα or mXinαCΔ as bait and two independent actin preys were grown on X-gal
containing selective plates for the same period of time. A stronger blue color was
developed from yeast cells containing mXinαCΔ bait and either actin prey, as compared
to the lighter color associated with yeast cells containing mXinα bait and either actin
prey. These results imply a stronger interaction between mXinαCΔ and actin than that
between mXinα and actin. (B) Results of β-galactosidase filter assay for the interaction
between mXinα and several mXinα-interacting proteins. After re-transformation of
mXinα bait and various preys into yeast cells, colonies grown on selective media were
transferred to membranes for X-gal assay. mXinα interacts with the extreme C-termini of
filamin b (aa#2,533-2,603). Additionally, mXinα interacts with gelsolin (aa#497-780),
and full-length constructs of tropomyosin, p120 catenin, and vinculin. However, mXinα
does not interact with full-length talin.
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Figure 2.10. Effect of GST-β-catenin on the binding of His-mXinα to actin filaments. An
actin (9.3 µM) binding assay was performed with increasing amounts of His-mXinα in
the absence (-) or presence (+) of GST-β-catenin (1.95 µM). After quantification of HismXinα in the pellet and supernatant fractions, data were plotted using SigmaPlot 9.0. The
presence of β-catenin appears to enhance the binding of mXinα to actin filaments. The
experiments were repeated three times and representative data are shown here.
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Figure 2.11. Actin bundle formation was accelerated by the presence of GST-β-catenin.
His-Xinα alone or mixed with an equal molar amount of GST-β-catenin (0.24 μM) was
incubated on ice for 2 hours. The proteins were then mixed with actin (2.4 μM) and
incubated at room temperature. At the time points indicated in the figure 10 μl aliquots
were taken and applied onto grids for negative staining. 50 electron micrographs were
randomly captured from each grid. The width of all the bundles in the micrograph were
measured and presented as histograms. Blue bar, without GST-β-catenin; red bar, with
GST-β-catenin. During the assay period, the number and size of bundles formed are timedependent. The bundles formed by mXinα in the presence of β-catenin appear to be more
and bigger than that formed by mXinα alone.
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Figure 2.12. Schematic model for how mXinα functions at the adherens junction. The
full-length mXinα molecule containing 15 Xin repeats exists in equilibrium between an
auto-inhibited state and open state, favoring the auto-inhibited state. β-catenin located at
the adherens junction region binds to the β-catenin-binding domain (β-catBD)
overlapping with the 12th-13th Xin repeats, and then the equilibrium shifts toward the
open state of the mXinα molecule, which facilitates subsequent binding/bundling of actin
filaments. Thus, mXinα is an integral component which links N-cadherin-mediated
adhesion to the actin cytoskeleton.
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-
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Figure 2.13. Characterization of actin bundles formed by His-mXinα at different molar
ratios of His-mXinα to actin. His-mXinα at different concentration was mixed with actin
(2.4 µM) and then the samples were processed to negative staining and electron
microscopy. The molar ratio of His-mXinα to actin in (A) equals to 2:1, (B) 1:25, (C)
1:100 and (D) 1:200. At the ratio 1:200, no bundles were found but some filaments were
tightly associated, as marked by the arrow (D). Bar=100 nm.
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Table 2.1. Computer assisted measurements of cell size and shape in transfected Chinese
Hamster Ovary (CHO) cells.
Transfected
Number
plasmid
of cells
pEGFP-C2a
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pEGFP-
Mean Area (µm2)
Mean
Roundness
perimeter (µm)
(%)
1,176.8 ± 676.8
172.4 ± 55.8
49.5 ± 16.1
115
1,162.7 ± 712.8
173.3 ± 50.7
47.1 ± 14.2
126
858.9 ± 538.7
159.1 ± 46.1
42.3 ± 16.2
p value: a vs. b
NS
NS
NS
a vs. c
0.0001
NS
0.0001
b vs. c
0.0001
0.03
0.004
mXinαb
pEGFPmXinαRΔ-3c
Note: Roundness is a percentage measure of how efficiently a given amount of perimeter
encloses the area. A circle has the largest possible area and has a roundness of 100%.
A t-test was performed to calculate p value if the samples passed the test of normality and
equal variance. Otherwise, a rank sum test was used. The designations a, b, and c denote
the three different constructs when comparing them statistically.
NS: not significant (p>0.05).
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Table 2.2. Scoring the population of transfected CHO cells showing GFP fused to mXinα
and to various deletion mutants associated with stress fibers.
Transfected
plasmid
pEGFP-mXinα
pEGFPmXinαRΔ-1
p valuea
pEGFPmXinαRΔ-2
p valuea
p valueb
pEGFPmXinαRΔ-3
p valuea
pEGFPmXinαCΔ
p valuea
p valuec
Number of
cells scored
934
516
% of total
Group I
Group II
Group III
Group IV
0.43
17.02
76.12
6.42
5.23
NS
88.57
<0.0001
4.07
<0.0001
2.13
NS
0.66
NS
NS
47.65
<0.01
<0.025
50.09
NS
<0.0001
1.60
NS
NS
95.78
<0.0001
4.22
<0.05
0
<0.0001
0
NS
0
NS
NS
11.02
NS
<0.0001
61.34
NS
NS
27.64
<0.025
<0.0001
1,066
1,043
1,107
Note: CHO cells transfected with GFP-full-length mXinα or various deletion derivatives
were scored under fluorescence microscope by counting numbers of detectable stress
fibers with GFP-signal per cell.
Transfected cells were then classified into group I with no detectable stress fiber, group II
with 3-10 stress fibers, group III with 10-20 stress fibers or Group IV with more than 20
stress fibers.
The Chi square test was performed to calculate p value.
NS, not significant difference; p valuea: pairwise comparisons between cells transfected
with plasmid for full-length mXinα and with plasmids for various deletion mutants; p
valueb: comparison between cells transfected with plasmid for mXinαRΔ-1 and with
plasmid for mXinαRΔ-2; p valuec: comparison between cells transfected with plasmid for
mXinαRΔ-2 and with plasmid for mXinαCΔ.
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CHAPTER III
ESSENTIAL ROLES OF AN INTERCALATED DISC PROTEIN,
mXINβ, IN POSTNATAL HEART GROWTH AND SURVIVAL
Preface
The data presented in the following chapter were included in a manuscript by
Qinchuan Wang, Jenny Li-Chun Lin, Benjamin E. Reinking, Han-Zhong Feng, Fu-Chi
Chan, Cheng-I. Lin, Jian-Ping Jin, Elisabeth A. Gustafson-Wagner, Thomas D. Scholz,
Baoli Yang, Jim Jung-Ching Lin, which was published in the Circulation Research
(Circulation Research. 2010; 106: 1468-1478). In this study, I tested the hypothesis that
mXinβ plays important roles in cardiac morphogenesis and function by generating and
characterizing the phenotypes of the mXinβ knockout mice. This study revealed that
mXinβ is essential for cardiac morphogenesis and function, as implicated by our previous
evolutionary study demonstrating that mXinβ is more conserved with the ancestral Xin. I
also showed that the underlying mechanism for the defects observed in mXinβ-knockout
hearts may involve dysregulation of multiple signaling pathways. Moreover, I observed a
failure in ICD maturation in the mXinβ-null hearts, which will be further investigated in
chapter IV.
In this study, I described the generation of mXinβ knockout mice. I showed that
loss of mXinβ leads to severe postnatal growth retardation and lethality before weaning.
Using histological techniques, I demonstrated that mXinβ-null animals have ventricular
septal defects (VSD), mis-organized myocardium and reduced heart weight. The mutant
hearts exhibit reduced proliferation and increased apoptosis, which may be responsible
for the morphological defects of the hearts. In collaboration with Dr. Thomas D. Scholz, I
obtained echocardiographic data to demonstrate the existence of diastolic dysfunction in
the mutant hearts, which is supported by the mis-organized myocardium and a delay in
the switching off of the slow skeletal troponin I. Importantly, through immunostaining of
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intercalated disc markers, I showed that mXinβ-null animals failed to form mature ICDs,
and a reduction of the small GTPase Rac1 activity might be responsible for this defect.
Finally, my data indicate that mXinβ functions in multiple signaling pathways,
suggesting that mXinβ might be an important scaffolding protein that organizes signaling
complexes at the ICDs.
Abstract
Rationale: The Xin repeat-containing proteins, mXinα and mXinβ, localize to the
intercalated disc (ICD) of mouse heart and are implicated in cardiac development and
function. The mXinα directly interacts with β-catenin, p120-catenin and actin filaments.
Ablation of mXinα results in adult late-onset cardiac cardiomyopathy with conduction
defects. An up-regulation of the mXinβ in mXinα-deficient hearts suggests a partial
compensation.
Objective: The essential roles of mXinβ in cardiac development and ICD
maturation were investigated.
Methods and Results: Ablation of mXinβ led to abnormal heart shape, ventricular
septal defects, severe growth retardation and postnatal lethality with no up-regulation of
the mXinα. Postnatal up-regulation of mXinβ in wild type hearts, as well as altered
apoptosis and proliferation in mXinβ-null hearts suggest that mXinβ is required for
postnatal heart remodeling. The mXinβ-null hearts exhibited a mis-organized
myocardium as detected by histological and electron microscopic studies, and an
impaired diastolic function as suggested by echocardiography and a delay in switching
off the slow skeletal troponin I. Loss of mXinβ resulted in the failure of forming mature
ICDs and the mis-localization of mXinα and N-cadherin. The mXinβ-null hearts showed
up-regulation of active Stat3 (signal transducer and activator of transcription 3) and
down-regulations of the activities of Rac1, IGF-1 (insulin-like growth factor 1) receptor,
Akt and Erk1/2 (extracellular-signal-regulated kinases 1/2).
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Conclusions: These findings identify not only an essential role of mXinβ in the
ICD maturation but also mechanisms of mXinβ modulating N-cadherin-mediated
adhesion signaling and its crosstalk signaling for postnatal heart growth and animal
survival.
Introduction
A regulatory network of transcription factors is known to control cardiac
morphogenesis. Although the core players in this network are highly conserved from
organisms with simple heart-like cells to those with complex four-chambered hearts, it
has been theorized and proven that expansion of this regulatory network by adding new
transcription factors is a major force for the heart to evolve new structures (Clark et al.,
2006; Olson, 2006). However, the addition of new transcription factors can only be a part
of the mechanism underlying the formation of complex hearts. The transcription factors
must act through their downstream targets, which are directly involved in cardiac
morphogenesis, growth and function. However, our inventory of such downstream targets
remains incomplete.
The Xin repeat-containing proteins from chicken and mouse hearts (cXin and
mXinα, respectively) were first identified as a target of the Nkx2.5-Mef2C pathway
(Barbato et al., 2005; Wang et al., 1999). Another mouse Xin protein, mXinβ (or
myomaxin), has been subsequently identified as a Mef2A downstream target (Huang et
al., 2006). Evolutionary studies suggest that Xin may be one of the factors that arose
when the heart evolved from simple heart-like cells to the complex true-chambered hearts
(Grosskurth et al., 2008). Functional studies reveal that Xin proteins are involved in heart
chamber formation and cardiac function in vertebrates (Barbato et al., 2005; Choi et al.,
2007; Wang et al., 1999). The striated muscle-specific Xin family of proteins are defined
by the presence of 15~28 copies of the conserved 16-amino acid (aa) Xin repeats, and
originated just prior to the emergence of lamprey, coinciding with the appearance of the
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true-chambered heart (Grosskurth et al., 2008). The Xin repeats are responsible for
binding actin filaments (Choi et al., 2007; Pacholsky et al., 2004; van der Ven et al.,
2006), whereas a highly conserved β-catenin binding domain (β-catBD) overlapping with
the Xin repeats is responsible for localizing Xin to the intercalated discs (Choi et al.,
2007; Grosskurth et al., 2008). Supporting the roles of Xin in heart chamber formation
and function, we have previously shown that knocking down the sole cXin in the chicken
embryo collapses the wall of heart chambers and leads to abnormal cardiac
morphogenesis (Wang et al., 1999).
In mammals, however, a pair of paralogous Xinα and Xinβ genes exists. Ablation
of the mouse mXinα gene does not affect heart development. Instead, the mXinα-deficient
mice show cardiac hypertrophy and cardiomyopathy with conduction defects during
adulthood. In the mXinα-deficient mice, mXinβ is up-regulated at both message and
protein levels, suggesting a compensatory role of mXinβ (Choi et al., 2007). Consistent
with this idea, both mXin proteins have highly conserved Xin repeats and β-catBD, as
well as other functionally undefined domains located in the N-terminals (Grosskurth et
al., 2008). On the other hand, the C-terminals of both proteins are more diverged,
suggesting that they also have distinct functions (Grosskurth et al., 2008). Since mXinβ is
more conserved than mXinα with the ancestral lamprey Xin that demarked the emergence
of heart chamber, we hypothesized that mXinβ might play an essential role in heart
morphogenesis. To test this hypothesis, we generated and characterized mXinβ knockout
mice. The mXinβ-null mice died prior to weaning and showed abnormal heart shape,
ventricular septal defects (VSDs), mis-organized myocardium and diastolic dysfunction.
The mechanisms underlying these cardiac defects involved dys-regulation of the Ncadherin-mediated signaling pathway and its crosstalks via abnormally activated Stat3
and depressed Rac1, IGF-1 receptor (IGF-1R), Akt and Erk1/2 activities.
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Materials and Methods
All animal procedures were approved and performed in accordance with
institutional guidelines. The mXinβ-null line has been backcrossed to and maintained in
C57BL/6J. All of phenotypes observed earlier remained the same.
5’ and 3’-RACE (rapid amplification of cDNA ends) of
mXinβ cDNAs
We have previously obtained several overlapping cDNA clones from a custommade cDNA library constructed from poly(A)+ RNAs prepared from adult mXinα-null
mouse hearts (Choi et al., 2007). The composite sequence of these pBKX cDNA clones
has 4,382 bp covering 9 bp of exon 1, exon 2~6 and partial exon 7. Since exon 7 was
predicted to be a very large exon from genomic sequence, we PCR amplified from the
mXinβ genomic β14 clone to construct our cDNA plasmid containing 9,985bp. The
Northern blot analysis revealed that mRNA size of mXinβ was ~12 kb (Barbato et al.,
2005), and a stop codon had not been reached in our composite cDNA sequence,
indicating that the full length mXinβ cDNA had not yet been obtained. Therefore, 5’ and
3’-RACE cloning were further performed using Marathon cDNA amplification kit
(Clontech) as previously described (Wang et al., 1999) to obtain additional cDNA
sequence information. For 5’-RACE, two antisense primers 5’TGCCTCCTGCTCAGCTCTGCTCTCATGTCG-3’ (nucleotide #463~434 relative to
mXinβ-A) 5’-GGGACAGCGCCTCCAGGAGATCCGACTG-3’ (nucleotide #251~224)
located in exon 5 and exon 3, respectively, were used in different experiments. For 3’RACE, primers 5’-ACACCACCTTCCCCACCAAGGAGTCGTTCA-3’ (nucleotide
#8,990 ~ 9,019), and 5’-CTTTGACTTCAAGCATGCCCCACCGACC-3’ (nucleotide
#9,808~9,835) located within the predicted exon 7, as well as 5’GGCCGCCTGAAGTGACCATCCCTGTTCC-3’ (nucleotide #10,515~10,542) in exon 9
were used. The 5’ and 3’ sequences of mXinβ messages obtained from RACE were
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submitted to GenBank (accession no. EU286528~286531). Accordingly, three different
full-length cDNAs were further constructed from the 9,985bp fragment and these RACE
products (Figure 3.1 A). To characterize these full-length cDNAs, the constructs were
subcloned into pcDNA1.1 (Invitrogen) and pEGFP-C2 (Clontech) vectors and transfected
to Chinese Hamster Ovary (CHO) cells. Immunofluorescence microscopy and Western
blot analysis were performed on the transfected cells to verify protein products.
Generation of mXinβ isoform-specific antibodies
To generate isoform-specific antibodies that recognize either mXinβ or mXinβ-a,
we subcloned the cDNA fragments encoding the isoform-specific regions of mXinβ
(encoding aa#3,255-3,278) and mXinβ-a (aa#3,256-3,300) (Figure 3.1B) into the
pGEX4-T2 vector (GE Healthcare Life Sciences). GST-fused recombinant proteins were
produced in BL21(DE3)pLysS bacteria and affinity purified with Glutathione Sepharose
4B beads (Amersham). The purified proteins were used to immunize rabbits to produce
mXinβ and mXinβ-a specific antisera (Cocalico Biologicals, Inc.). To affinity-purify
antibodies specific to mXinβ (U1040) and mXinβ-a (U1043), the antisera were first
passed through GST-conjugated Sepharose 4B column to deplete the anti-GST
antibodies. U1040 and U1043 were then affinity-purified with their respective antigenconjugated Sepharose 4B column.
Construction of mXinβ targeting vector and generation of
mXinβ-null mice
Using previously cloned pBKX-2 cDNA (accession no. AY775570) as a probe,
we screened λfix II genomic library constructed from mouse strain 129SVJ genomic
DNA (Stratagene) and obtained 8 overlapping clones including β14 containing portion of
intron 3 to exon 8 of the mXinβ gene. Subclones of β14 genomic fragments were used to
construct targeting vector for inactivating mXinβ gene by replacing portions of exon 6,
intron 6 and portion of exon 7 with a LacZ-Neor cassette, since these regions encode
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highly conserved DNA-binding domain (DBD), β-catenin-binding domain (β-catBD) and
Xin repeats (Figure 3.1 B). The linearized targeting vector was electroporated into R1
embryonic stem (ES) cells at the University of Iowa Gene Targeting Facility. After
selection, G418-resistant ES clones were screened for the presence of the targeted locus
by Southern blot analysis (Wang et al., 1999). Positive clones were expanded and
microinjected into C57BL6 blastocysts to generate chimera. Chimeric male mice were
analyzed for germ-line transmission by Southern blot analysis and PCR genotyping. For
PCR genotyping, genomic DNA extracted from the toes or tails of mice was used. The
primer pairs used were 5’-GACAGGCTGGCCATACTCAA-3’ and 5’ACATTTTTCTAAGGCTTTTCTCAA-3’ for the endogenous mXinβ locus, and 5’CCTGGCCCCTACTCCTACCTTTTT-3’ and 5’-CGGGCCTCTTCGCTATTACG-3’ for
the targeted locus. The heterozygous mice were back-crossed to C57BL6 for at least 7
generations, maintained in C57BL6 background, and used for obtaining most results
reported here. All phenotypes observed in mXinβ-null mice earlier in a mixed
C57BL6/129SVJ background essentially remained the same in the mutants with C57BL6
background.
Histological staining, immunofluorescence, and assessment
of ventricular myoarchitecture
Hearts excised from wild-type, heterozygous and homozygous littermates were
fixed in 10% formalin in phosphate buffered saline (PBS) for one to two days at 4 °C.
Tissue processing, Hematoxylin and Eosin (H&E) staining, and Masson’s trichrome
staining were carried out as previously described (Choi et al., 2007). For
immunofluorescence microscopy, a pair of hearts from both wild type and mXinβ-null
littermates were arranged in the same orientation and embedded in Tissue-Tek O.C.T.
Compound (Sakura Finetek USA, Inc.) in the same Peel-A-Way mold (Thermo
Scientific) and frozen in liquid nitrogen for 30 seconds. Frozen sections were cut at 4 µm
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thickness and mounted onto Superfrost PLUS slides (Thermo Scientific). Subsequent
immunofluorescence microscopy was carried out as previously described (Choi et al.,
2007). The primary antibodies used included rabbit polyclonal U1697 anti-mXinα (Choi
et al., 2007), U1040 anti-C-terminus of mXinβ, anti-pan cadherin (Sigma), mouse
monoclonal anti-β-catenin (Zymed Laboratories Inc.), anti-p120-catenin (Zymed), antiN-cadherin (Zymed), anti-connexin 43 (Cx43) (Chemicon International, Inc.), CT3 anticardiac troponin T (cTnT) (Warren and Lin, 1993), as well as rat monoclonal LAM-1
anti-laminin (ICN Biochemical, Inc.).
For measuring ventricular myoarchitecture, ventricles from P0.5 wild type and
mXinβ-null mice were processed according to the method described previously (Ishiwata
et al., 2003). The area of the myocardial layer was measured at the lateral free wall in
both left and right ventricles using sections obtained at the levels of major papillary
muscles. The compact and trabecular regions were calculated as areas per 1 µm
myocardium as defined previously (Ishiwata et al., 2003).
Transmission electron microscopy
P15.5 littermates were anesthetized and dissected. The major blood vessels
entering and exiting the heart were clamped with a hemostat, and the right atrium was cut
open for drainage of blood. Using an AutoMate In Vivo manual gravity perfusion system
(Braintree Scientific, Inc.), the heart was infused from the left ventricle with 2 ml of
Locke’s solution (146 mM NaCl, 5.63 mM KCl, 2.38 mM NaHCO3, 1.63 mM CaCl2,
0.1% lidocaine and 50 USP units/ml Heparin, pH7.4), and then perfusion-fixed with 5 ml
of primary fixative (2.5% glutaraldehyde, 4% paraformaldehyde, 30 mM CaCl2, 0.1 M
sodium cacodylate, pH7.4). After perfusion-fixation, the left ventricle of dissected heart
was diced into small blocks and kept in fixative for overnight followed by sequential
treatments with osmium tetraoxide and uranyl acetate. The tissue blocks were then
processed and embedded in epon resin. Ultrathin sections were cut, mounted, post-stained
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and observed under a JEOL1230 electron microscope (Central Microscopy Research
Facility, University of Iowa).
Proliferation and apoptosis
To study proliferation, littermates were intraperitoneally injected with 50µg
bromodeoxyuridine (BrdU, Invitrogen) per gram of body weight. Four hours later, hearts
were dissected from the injected mice and processed for frozen sections. At least five
sections that cut through both mitral and tricuspid valves were collected from each heart.
The sections were fixed in methanol for 20 minutes, denatured with 2N HCl for 2 hours
and neutralized with 0.1 M Sodium Borate. Immunofluorescence staining was then
performed on the neutralized sections with mouse monoclonal anti-BrdU (Developmental
Studies Hybridoma Bank, University of Iowa). Nuclei were counter-stained with 4’,6’diamidino-2-phenylindole (DAPI) as described previously (Grosskurth et al., 2008). To
quantify the frequency of BrdU-positive nuclei in the sections, 5 fluorescence images
were taken with a 10x objective from comparable regions of each section. The number of
BrdU positive nuclei and total nuclei were counted with the analyze particle function of
ImageJ (http://rsbweb.nih.gov/ij/). Student’s t-test was used to compare the frequency of
BrdU-positive cells between wild type and mXinβ-null hearts. For each stage, at least
three wild type and three mXinβ-null hearts were used.
To study apoptosis, immunofluorescence staining with ab13847 anti-active
caspase 3 antibody (Abcam, Inc.) was performed on frozen sections of wild type and
mXinβ-null hearts. The cardiomyocytes and nuclei were counter-stained with CT3 anticTnT and DAPI, respectively. Image collection and analysis were performed as described
above.
Cardiomyocyte width measurement
Frozen sections of postnatal day 3.5 (P3.5) and P12.5 wild type and mXinβ-null
hearts were immunostained with anti-laminin to outline cardiomyocytes, anti-cTnT to
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label cardiomyocyte cytoplasm and DAPI to label nuclei. Images were taken from
comparable area of the heart sections of each genotype. To quantify the width of
cardiomyocytes, cross sections of cardiomyocytes that cut through the nuclei were
identified. Then the length of the shorter axis that ran through the nuclear center of the
cross-sectioned cardiomyocytes was measured with Openlab 4.03 (Improvision Inc.).
Body weight, heart weight and liver weight measurements
Body weight was measured just before the mice were dissected. For heart and
liver weights, heart and liver were dissected out from mice, rinsed in cold PBS or Tris
buffered saline (TBS, pH7.5), blotted dry and then weighed.
Echocardiography
The mice were anesthetized and placed on a warming platform, and an
appropriately sized nose cone was placed over the pup nose. Anesthesia was maintained
at a minimum to suppress spontaneous movements. Heart rate was maintained between
350 and 600 beats per minute. Temperature was monitored with a rectal thermometer and
maintained between 35 and 36 oC.
Echocardiograms were performed using the Visual Sonics Vevo 770 High
Resolution Imaging System and software (Visual Sonics, Inc.), as described (Hinton et
al., 2008). The 704 (center frequency 40 MHz, focal length 10 mm), 707B (center
frequency 30 MHz, focal length 12.7 mm), and 710B (center frequency 25 MHz, focal
length 15 mm) RMV scan heads were used. Scan heads were interchanged during each
study to allow for optimal image acquisition. Parasternal long axis, parasternal short axis
and apical four chamber views were obtained in all animals. Pulsed wave Doppler
recordings were made across the left ventricular outflow tract, right ventricular outflow
tract and mitral valve when the Doppler sample volume was thought to be parallel to
flow. Doppler tracings could not be obtained in every animal and those that were more
than 20 degrees from parallel were not used for data analysis. M-mode recordings were
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obtained of the right and left ventricles in the parasternal short axis view at the level of
the left ventricular papillary muscles. Measurements were made of the interventricular
septum thickness in diastole and systole (IVSd, IVSs), left ventricular internal dimension
in diastole and systole (LVIDd, LVIDs), and left ventricular posterior wall thickness in
diastole and systole (LVPWd, LVPWs). These measurements were then used to calculate
the left ventricular ejection fraction (EF) and fraction shortening (FS), left ventricular
diastolic and systolic volumes (LVVd, LVVs), and left ventricular mass. Calculations
were made by the Vevo 770 software. Measurements were made in accordance with the
American Society for Echocardiography Guidelines (Lang et al., 2006).
Western blot analysis
For Western blot, tissues were dissected from anesthetized mice, rinsed in cold
TBS, blotted dry, weighed and then homogenized by a Pro200 homogenizer with MultiGen 7 generators (Pro Scientific Inc.) in buffer (40 µl buffer per mg heart weight)
containing 20 mM HEPES (pH 7.2), 25 mM NaCl, 2 mM EGTA, 2 mM Na3VO4, 25 mM
β-glycerophosphate, 50 mM NaF, 1% Triton X-100 and complete protease inhibitor
cocktail (cat#11836145001, Roche Applied Sciences). An equal volume of 2X SDSPAGE gel sample buffer was added to the homogenate. After heating at 100 °C for 5
minutes, the homogenate was stored at -70 °C as aliquots until use. Western blot analysis
was carried out as described previously (Choi et al., 2007). The primary antibodies used
included rabbit polyclonal U1013 anti-mXin (Choi et al., 2007; Sinn et al., 2002) and
anti-α-actinin (a generous gift from Dr. K. Burridge, UNC Chapel Hill); mouse
monoclonal CH1 anti-α-tropomyosin,(Lin et al., 1985) CT3 anti-cTnT (Warren and Lin,
1993), TnI-1 anti-troponin I (TnI) (Barbato et al., 2005), FA2 anti-α-myosin heavy chain
(MHC) (Choi et al., 2007), anti-β-MHC (Sigma), anti-plakoglobin (BD Biosciences),
ARC03 anti-Rac1 (Cytoskeleton, Inc.), and ARH03 anti-RhoA (Cytoskeleton, Inc.); as
well as rabbit monoclonal antibodies (Cell Signaling Technology, Inc.) including C67E7
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anti-pan Akt , D9E anti-p-Akt(S473), C31E5E anti-p-Akt(T308), DA7A8 anti-p-IGF-1R,
D3A7 anti-p-Stat3(Y705), 79D7 anti-Stat3, and C80C3 anti-p-Jak2(Y1007) and rabbit
polyclonal antibodies (Cell Signaling Technology, Inc.) including anti-p-GSK3β(S9) and
anti-p-Erk1/2.
Analysis of immunolocalization of N-cadherin and Cx43
during postnatal heart development
Frozen 4-µm sections of hearts from wild type and mXinβ-null littermates were
processed for immunofluorescence labeling with anti-N-cadherin or anti-Cx43 as
described above. The proportion of label at the myocyte termini during postnatal
development was determined by the previously described method (Angst et al., 1997;
Choi et al., 2007; Peters et al., 1994) with a slight modification. Briefly, micrographs
were taken from comparable regions of the ventricular myocardia and analyzed in
ImageJ. To facilitate analysis, images of individual cardiomyocytes that had been
longitudinally sectioned were extracted from the micrographs. Differential interference
contrast micrographs were used to aid this process. The termini of individual
cardiomyocytes were defined by making a rectangular box at both ends of the
cardiomyocytes. The width of each box was 10% of the longitudinal axis of the
cardiomyocyte. The percentage of terminally localized N-cadherin or Cx43 was
calculated by dividing the number of fluorescence pixels in the termini with the number
of fluorescence pixels in the whole cardiomyocytes.
Rac1 and RhoA activity assay
Active Rac1 and RhoA pull-down assays were carried out essentially as described
(Noren et al., 2000) with slight modifications. For active Rac1 pull-down assay, P7.5
mouse heart of each genotype was quickly dissected, rinsed in cold TBS, weighed and
then homogenized in ice cold Rac1 lysis buffer (40 µl buffer per mg heart weight)
containing 50 mM Tris-HCl (pH7.6), 150 mM NaCl, 1% TritonX-100, 10 mM MgCl2, 10
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µg/ml Aprotinin, 10 µg/ml Leupeptin, and 1 mM phenylmethyl sulfonyl fluoride
(PMSF). The homogenate was cleared by centrifugation at 20,000 x g for 15 minutes.
Then, 200 µl Rac1 lysis buffer containing 100 µg GST-Pak PBD (GST-Pak pre-bound to
Glutathione sepharose) beads were added to 400 µl cleared homogenate to pull-down
active Rac1. After washing the beads, GTP-bound (active) Rac1 was recovered in SDSPAGE gel sample buffer, followed by Western blot analysis.
For active RhoA pull-down assay, the heart homogenate was prepared as
described for the Rac1 activity assay, except that the lysis buffer for RhoA assay
contained 50 mM Tris-HCl (pH7.2), 500 mM NaCl, 10 mM MgCl2, 1% TritonX-100,
0.5% Sodium Deoxycholate, 0.1% SDS, 10 µg/ml Aprotinin, 10 µg/ml Leupeptin, and 1
mM PMSF. For 400 µl cleared homogenates, 50 µg of GST-Rhotekin RBD (GSTRhotekin pre-bound to Glutathione sepharose) beads were used to pull-down GTP-bound
RhoA. After washing the beads, active RhoA was recovered in gel sample buffer,
followed by Western blot analysis.
Results
Generation of mXinβ-null mice
To construct a targeting vector, we cloned full-length mXinβ cDNAs and the
corresponding genomic fragments. Alignment of these sequences revealed that the mXinβ
gene contains nine exons and encodes three mRNA species (mXinβ-A, mXinβ-B and
mXinβ-C) in adult heart through alternative splicing of exon 8 and alternative usage of
polyA signals (Figure 3.1 A). Both mXinβ-A and mXinβ-B encode a polypeptide of 3,283
aa residues (termed mXinβ), whereas mXinβ-C is predicted to encode a protein of 3,300
residues (termed mXinβ-a) (Figure 3.1 B). By sequencing 24 randomly picked
transformants generated from 3’-RACE, we found 23 clones representing either mXinβ-A
or mXinβ-B, suggesting that mXinβ is the major isoform. Force-expression of the cloned
mXinβ-B cDNAs in CHO cells confirmed that mXinβ-B encoded the protein having the
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same size as endogenous mXinβ and reacting to anti-mXin antibody (Figure 3.1 C).
Furthermore, force-expressed mXinβ co-localized with actin filaments to stress fibers and
cell cortex (Figure 3.1 D).
Using multiple tissue Western blot, mXinβ was detected only in the striated
muscles such as tongue, heart and diaphragm (Figure 3.2 A). During postnatal heart
development, the expression of mXinβ increased at least 3 folds from postnatal day 0.5
(P0.5) to P13.5 (Figure 3.2 C). The timing of this up-regulation of mXinβ coincides with
the period for intercalated disc maturation (Perriard et al., 2003; Sinn et al., 2002).
A targeting vector was designed to delete the genomic region that encodes the
highly conserved β-catBD and Xin repeats (Figure 3.3 A). After electroporation and
selection, resistant embryonic stem (ES) clones were screened by Southern blot analysis
(Figure 3.3 B). The positive ES clone was used to generate chimeric founders. After
confirming germ-line transmission, the heterozygous progeny were further crossed to
obtain mXinβ-null mice. The genotypes of the resulting littermates were determined with
tail DNAs by Southern blot and by PCR genotyping (Figure 3.3 C).
All mXinβ-null mice die before weaning
Northern blot analysis revealed a complete loss of mXinβ message in
homozygotes and a reduction in heterozygotes (Figure 3.3 D). Western blot analyses with
antibody U1013 (Choi et al., 2007) recognizing both mXinα and mXinβ (Figure 3.3 E,
top blot) or with antibody U1040 recognizing C-terminal of mXinβ (data not shown)
verified a complete loss of mXinβ in homozygotes and a reduced level in heterozygotes.
The mXinβ-null hearts expressed similar amounts of mXinα-a, mXinα, α-actinin and αtropomyosin (α-TM) as their age-matched counterparts (Figure 3.3 E).
At birth (P0.5), the number of the mXinβ-null pups from heterozygous crosses
was smaller than the expected number (Table 3.1), however, this reduction was not
statistically significant (p=0.17, Chi square test). In contrast, from P3.5 and on, the
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number of viable mXinβ-/- mice was significantly lower than the expected one. No viable
mXinβ-/- mice could be observed at weaning stage. Thus, these observations suggest that
mXinβ is essential for postnatal mouse survival.
Loss of mXinβ leads to severe growth retardation
The mXinβ-null mice had severely retarded growth and reduced activity. The skin
of newborn mXinβ-/- mice was apparently paler than their littermates, suggesting a
systemic circulation defect. Great vessels in the newborn mXinβ-null mice were normal
(Figure 3.4). The body weight (BW) of P0.5 mXinβ-/- mice was about 14.3% lighter than
wild type or heterozygous littermates (Figure 3.5 A). From birth to P12.5, the mXinβ-null
mice also gained weight more slowly than their littermates (Figure 3.5 A). At P12.5,
mXinβ-null mice weighed only about 45% of wild type or heterozygous mice. The loss of
just one copy of mXinβ in heterozygotes had neither effect on BW nor on viability.
Neonatal mXinβ-/- pups apparently breathed normally, and milk was always visible in
their stomach, suggesting that a weakness in skeletal muscles is unlikely to be the major
cause for the growth retardation and lethality.
The heart weights (HW) of newborn wild type and mXinβ-null pups were similar.
However, from P3.5 to P12.5, the wild type hearts grew much faster than mXinβ-null
hearts. As a result, both P7.5 and P12.5 wild type hearts were significantly larger than
mXinβ-null counterparts (Figure 3.5 B), suggesting that mXinβ is required for postnatal
heart growth. Similar to its effects on BW, the loss of one copy of mXinβ in
heterozygotes did not affect their heart size (data not shown). The HW/BW ratio of
mXinβ-/- mice at most of postnatal stage except P3.5 was significantly higher than that of
wild type mice (Figure 3.5 C) due to significantly smaller BW in mXinβ-null mice.
Similar to the hearts, mXinβ-non-expressing organ such as liver (Figure 3.5 D) of mXinβnull mice also became significantly smaller between P3.5 and P7.5. However, the liver
weight (LW) to BW (LW/BW) ratios of wild type and mXinβ-null mice remained no
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difference (Figure 3.5 E). Thus, the loss of mXinβ affected the mXinβ-expressing and
mXinβ-non-expressing organs differently.
Loss of mXinβ results in VSDs, abnormal heart shape and
mis-organized myocardium
About 15% (5/33) of newborn mXinβ-null hearts had abnormal shape (Figure 3.6
B). VSD was detected in 58% (7/12) of newborn mXinβ-null hearts analyzed by serial
section analysis (arrow in Figure 3.6 D). The VSD could be found in any locations within
the muscular septum, and could be small, large or multiple. However, the VSD could not
be the cause of postnatal lethality, since 42% of mXinβ-null mice without VSD also
became small and weak, and died before weaning. Mis-organized myocardium (noncompaction in right ventricle) could be detected in mXinβ-null hearts as early as
embryonic day 14.5 (E14.5) (Figure 3.6 F&F’). Thus, mXinβ-null embryo may already
have a defect in heart function, leading to a slight but significant reduction in BW at birth
(Figure 3.5 A). However, this defect may not be enough to cause embryonic lethality,
since no significant loss of newborn mXinβ-null mice was found (Table 3.1).
Furthermore, all mXinβ-null neonatal hearts examined showed various degree of misorganized myofibers within myocardium (an example shown in Figure 3.6 H’&H’’).
Electron microscopic (EM) analysis of P15.5 mXinβ-null hearts detected no sarcomere
disorganization within each myocytes (Figure 3. 6J), suggesting no myofilament disarray
in mutant hearts.
Developing mXinβ-null hearts exhibit diastolic dysfunction
Since all mXinβ-/- mice exhibited mis-organized myocardium, we next analyzed
chamber size, wall thickness and cardiac function by echocardiography. Because
echocardiographic results from wild type and heterozygotes were very similar, we treated
them as a control group for the comparison to mXinβ-null group (Table 3.2). We
observed a reduction in left ventricular internal dimension and volume of both P3.5 and
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P12.5 mXinβ-null hearts during diastole and systole (Table 3.2). In contrast, there was no
difference between control and mXinβ-null mice in heart rate, left ventricular posterior
wall thickness and interventricular septum thickness (Table 3.2). Furthermore, the
mXinβ-null hearts had normal or slightly higher systolic function, as determined by the
ejection fraction and the fraction shortening (Table 3.2). Using pulsed wave Doppler
recordings, we found that mXinβ-null hearts exhibited abnormal ventricular filling. In
mXinβ-/- mice, the mitral inflow E-wave (early filling) but not A-wave (atrial
contraction) peak velocity was reduced (Figure 3.7), and the E/A ratios were also
significantly smaller (Table 3.2). These results suggest a diastolic dysfunction in mutant
hearts as early as P3.5. However, this diastolic dysfunction was not due to increased
fibrosis that could stiffen the myocardium, because Trichrome staining detected no
increase in fibrosis at P11.5 (Figure 3.8).
Developmental changes in ventricular diastolic function correlate well with
changes in myoarchitecture (compact versus trabecular areas in ventricles) (Ishiwata et
al., 2003). In general, the peak E-wave velocity is exponentially correlated with the area
of compact region per unit myocardium, whereas the peak A-wave velocity is correlated
with the area of trabecular region per unit myocardium. Using similar measurement in
newborn mXinβ-null mice, we found a significant reduction in the area of left ventricle
compact myocardium and a trend of increase in the area of left ventricle trabecular
myocardium in mutant hearts (Table 3.3). Similar trends of decrease in compact area and
increase in trabecular area were also observed for right ventricle (Table 3.3). These
results again support diastolic dysfunction associated with newborn mXinβ-null hearts.
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The delay in switching off slow skeletal troponin I (ssTnI)
also supports diastolic dysfunction associated with mXinβnull mice
Apparent preservation of systolic function and presence of diastolic dysfunction
in mXinβ-null hearts led us to examine the expression levels and isoform switches of
contractile and regulatory proteins. The observations of normal expression levels of αactinin and α-TM (Figure 3.3 E) as well as normal timing of switching from β-myosin
heavy chain (MHC) to α-MHC (Figure 3.9) and from embryonic cardiac troponin T
(ecTnT) to adult cTnT (acTnT) (Figure 3.10 C) largely support that mXinβ-null hearts
having normal systolic function. In contrast, a significant delay in switching off ssTnI
was detected in P7.5 and P13.5 mXinβ-null hearts (Figure 3.10 B). This delay may allow
mutant hearts to gain increased Ca2+-activated myofilament tension to compensate for
function, because ssTnI has higher Ca2+ sensitivity than cTnI, which is supported by
previous study comparing force generations between ssTnI- and cardiac troponin I
(cTnI)-expressing cardiomyocytes (Westfall and Metzger, 2001). Nonetheless, transgenic
mice ectopically expressing ssTnI in the heart exhibit impairments of cardiomyocyte
relaxation and diastolic function (Fentzke et al., 1999). Together, the delay in switching
off ssTnI also supports diastolic dysfunction in mXinβ-null heart. It should be noted that
mXinβ-null hearts did not up-regulate N-terminal truncated cTnI (cTnI-ND) (Figure 3.10
B), which has been previously shown to enhance ventricular diastolic function in
transgenic mice (Barbato et al., 2005).
Developing mXinβ-null hearts exhibit an increased
apoptosis as well as a decreased proliferation
Apoptosis and proliferation contribute greatly to myocardial remodeling during
postnatal development (Fernandez et al., 2001). Thus, we asked whether defects in these
processes might contribute to the mis-organization of mutant myocardium. The wild type
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hearts had high apoptosis only at P0.5, as detected by anti-active caspase 3, which then
rapidly declined to a minimal level at P7.5, similar to that of adult heart (Fernandez et al.,
2001) (Figure 3.11 A, A’ and E). In contrast, the level of apoptosis in P0.5 mXinβ-null
hearts decreased more slowly and remained significantly higher at P3.5 and P7.5 (Figure
3.11 B, B’ and E). Using bromodeoxyuridine (BrdU) labeling, we found that there was
no difference in proliferation rate in mXinβ-null and control hearts until P7.5, at which
mutant hearts showed slightly reduced cell proliferation (Figure 3.11 C, D and F).
Therefore, slightly decreased proliferation and increased apoptosis in mXinβ-null hearts
postnatally may in part account for the smaller HW and the mis-organized myocardium.
Cardiomyocyte organization was compared from cross-sections of individual
cardiomyocytes of similar regions of littermate hearts. The cTnT-positive cardiomyocytes
were outlined by anti-laminin for shape and width comparison. At P3.5, there was no
detectable difference between wild type and mXinβ-null cardiomyocytes in either cell
shape or cell width (Figure 3.11 A, B and E). In contrast, by P12.5, mXinβ-/cardiomyocytes became more irregularly shaped (Figure 3.12 D) and smaller in cell
width (Figure 3.12 E).
mXinβ-null hearts fail to develop mature intercalated discs
At the first two weeks of age, mXinα, N-cadherin and β-catenin progressively
coalesce to the termini of aligned cardiomyocytes to form mature intercalated discs
(Angst et al., 1997; Sinn et al., 2002). We asked whether mXinβ plays a role in the
intercalated disc maturation. In P16.5 wild type hearts, majority of mXinβ, N-cadherin
and mXinα (Figure 3.13 A, C and E) as well as β-catenin and p120-catenin (data not
shown) were already localized to the mature intercalated discs. In contrast, most Ncadherin (Figure 3.13 D) and β-catenin (data not shown) found in the P16.5 mXinβ-null
hearts remained as small puncta along the lateral contacts of cardiomyocytes, while p120catenin (data not shown) and mXinα (Figure 3.13 F) puncta became dispersed throughout
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the cardiomyocytes. These results suggest that mXinβ is essential for promoting and
maintaining the localization of adherens junctional components and mXinα to the mature
intercalated discs.
The wild type and mXinβ-null mice from newborn to 2~3 weeks of age appeared
to express comparable amounts of N-cadherin and connexin 43 (Cx43) (Figure 3.9,
Figure 3.14 A, and some data not shown). Both N-cadherin and Cx43 continued to
accumulate to the myocyte termini of the wild type mice from P15.5 to P18.5, whereas
the terminal distribution of both molecules remained unchanged in mutants (Figure 3.14
B), again suggesting a defect in the maturation of intercalated discs. EM analysis
revealed that the developing intercalated disc at the cell termini of P15.5 mXinβ-null
hearts was smaller than the wild type counterparts (arrows in Figure 3.13 G & H). At
higher magnification, the membranes at the maturing intercalated disc of mXinβ-null
cardiomyocytes were less convoluted and less wavy (Figure 3.13 J), suggesting a
depressed membrane activity at the termini of mXinβ-null cells. At the lateral membrane
contacts, developing T-tubules could be detected in both mutant and control cells (* in
Figure 3.13 G and H), and less difference in the membrane activity was observed.
The mXinβ-null hearts increased Stat3 activity but
decreased Rac1, IGF-1R, Akt and Erk1/2 activities
Accumulated lines of evidence suggest that N-cadherin-mediated adhesion
signaling is critical for intercalated disc integrity and cardiac function (Fukuyama et al.,
2006). Cadherin and its associated catenins are also known to interact with many
signaling molecules, providing the ability to cross-talk with other signaling pathways
such as receptor tyrosine kinase-, cytokine receptor- and G protein coupled receptormediated signaling. We asked whether impairing intercalated disc maturation by the loss
of mXinβ could lead to abnormal activities of Rho GTPase, Stat, Akt and Erk, important
effectors in relaying signaling for postnatal heart development.
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Using GST-Pak PBD and GST-Rhotekin RBD beads to pull-down active forms of
Rac1 and RhoA, respectively, we found that relative GTP-bound Rac1 in P7.5 mXinβnull hearts was reduced to ~65% of the control, whereas the active RhoA level in mXinβnull hearts did not change significantly (Figure 3.15 A). A reduction of Rac1 activity may
result in less dynamic membranes at the termini of mutant cells, which was indeed
suggested by the EM observation. Using phospho-specific antibodies to assess the
activation of key signaling molecules involved in proliferation, growth and survival
(Figure 3.15 B), we found an increased Stat3 activity, as suggested by increased level
(Figure 3.15 C) and nuclear localization (data not shown) of p-Stat3(Y705) (tyrosinephosphorylated Stat3 at #705), persistently in mXinβ-null hearts starting from P0.5. This
Stat3 activation was not correlated to the activation of Jak2 (Janus kinase 2) (one member
of non-receptor tyrosine-protein kinases upstream of Stat3) (Figure 3.15 C), suggesting
that other Jaks and/or c-Src may be involved in the activation of Stat3. Alternatively,
defects in negative regulators of Stat3, such as suppressor of cytokine signaling 3
(SOCS3) or tyrosine phosphatases, may participate in the abnormal activation of Stat3 in
mutant hearts. Moreover, the activations/phosphorylations of Akt, GSK3β (glycogen
synthase kinase 3β, a downstream target of Akt), Erk1/2 and IGF-1R were significantly
depressed in mutant hearts starting from P7.5, whereas the total proteins of Akt and Grb2
(growth factor receptor-bound protein 2) in mutant and control hearts remained the same
(Figure 3.15 C). The persistent activation of Stat3, although not 100% penetrant, precedes
the reductions in the activations of growth-related signaling molecules.
Discussion
In this study, we demonstrate that an intercalated disc-associated and Xin repeatcontaining protein, mXinβ, is required for postnatal heart development. First, the
postnatal up-regulation of mXinβ coincides with the maturations of the intercalated disc
(Perriard et al., 2003; Sinn et al., 2002), T-tubule and sarcoplasmic reticulum (Hirakow
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and Gotoh, 1980), as well as diastolic function (Zhou et al., 2003). Second, ablation of
mXinβ leads to abnormal heart shape, VSD, diastolic dysfunction, severe growth
retardation, and postnatal lethality. Third, loss of mXinβ results in failure of forming
mature intercalated disc. Our data further identify that the proper clustering of N-cadherin
to form intercalated disc regulates the Stat3 activity and activates the Rac1, IGF-1R, Akt
and Erk1/2 activities, which are required for postnatal heart growth/hypertrophy (Clerk
et al., 2001; Satoh et al., 2006; Yamane et al., 2007).
How does the intercalated disc mature?
Postnatal maturation of intercalated discs is characterized by gradual clustering of
N-cadherin complexes/puncta from lateral localization to termini of aligned
cardiomyocytes. Such a clustering process likely involves modulating the interaction
between cadherins and underlining actin cytoskeleton. In a classic view, the actin
bundling protein, α-catenin, binds β-catenin to organize the adhesion complex that links
to actin cytoskeleton (Pokutta and Weis, 2002). However, this stable linkage role for αcatenin has not been proven; instead, compelling evidence suggests α-catenin being a
molecular switch that modulates actin cytoskeleton (Drees et al., 2005). Consistent with
this notion, two types of cadherin-mediated intercellular contacts are recently detected in
the adherens junctions of epithelia: a mobile and α-catenin-dependent contact associated
with a dynamic actin network as well as a stable and α-catenin-independent contact
associated with a stable actin patch (Cavey et al., 2008). The existence of this stable
contact suggests that an unidentified protein X has to link the cadherin/catenin complex
to actin patches. In the heart, the role of this unidentified protein X may be served by the
Xin repeat-containing proteins. We propose that developmental up-regulation and
functional hierarchy of mXinβ initiate the formation of mature intercalated discs. The
mXinα further reinforces the stability of intercalated discs. In support of this role, loss of
mXinβ leads to failure of forming mature intercalated discs and mis-localizations of
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mXinα and N-cadherin. On the other hand, mature intercalated discs form normally in the
mXinα-null heart (Figure 3.16), but eventually lose close membrane apposition between
cardiomyocytes at young adult. This structural defect progressively worsens by older age
(Choi et al., 2007).
Diastolic dysfunction may be responsible for heart failure
and lethality in mXinβ-null mice
The mXinβ-null hearts have normal systolic function and heart rate, but exhibit a
significant delay in switching off ssTnI and significant reductions in mitral early filling
(E-wave) peak velocity and E/A ratio, suggesting diastolic dysfunction. Impaired
diastolic function was also suggested by the left ventricle internal dimension and left
ventricle volumes being smaller in mXinβ-null mice. The detection of a significant
reduction in the compact areas of ventricles in newborn mutant hearts (Table 3.2I),
further supported a reduction in E-wave velocity (Ishiwata et al., 2003). The diastolic
dysfunction would lead to diminished cardiac output (stroke volume x heart rate) of
mutant hearts and could contribute in part to heart failure and postnatal lethality. The
mXinβ-null cardiomyocytes after P15.5 exhibited a significant reduction in the terminal
Cx43 localization (Figure 3.14), which may cause arrhythmic sudden death. However,
this spatial Cx43 alteration cannot be the cause for the loss of mXinβ-null mice at earlier
age (Table 3.1).
mXinβ regulates postnatal cardiac growth
In the heart, the Rac1 activation is essential for rearranging cytoskeleton to align
cardiomyocytes (Yamane et al., 2007), and for regulating mitogen-activated protein
kinases (Clerk et al., 2001) and NADPH oxidase activity (Satoh et al., 2006) for cardiac
hypertrophy. Moreover, transgenic mice expressing constitutively active Rac1 in the
heart develop dilated myocardium with high postnatal mortality (Sussman et al., 2000).
Most transgenic mice die within 2~3 weeks after birth, suggesting that postnatal heart
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development requires an intricate regulation of Rac1 activity. It is also known that classic
cadherin engagement activates Rac1 through c-Src-PI3K-Vav2, and Vav2 is a guanine
nucleotide exchange factor capable of binding to p120-catenin (Fukuyama et al., 2006;
Noren et al., 2000). The loss of mXinβ may dys-regulate this signaling, leading to a
down-regulation of Rac1 activity and forming less convoluted, less wavy, and less stable
intercalated discs. The loss of mXinβ may also dys-regulate cytokine/AngII/growth
hormone-mediated signaling, leading to a persistent activation of Stat3 (Figure 3.17). The
activation of Stat can promote IGF-1 production (Honsho et al., 2009), which would
facilitate postnatal heart growth. However, the lack of mature intercalated discs in mutant
hearts reduced the activities of IGF-1R, Akt and Erk1/2, resulting in severely retarded
growth.
In summary, we have identified that mXinβ, as a critical component for the
intercalated disc maturation, is essential for postnatal heart development. Our findings
provide the first insights into its function of transducing the N-cadherin-mediated
adhesion and crosstalk signalings by regulating the activities of Stat3, Rac1, Erk1/2 and
Akt. Ablation of mXinβ leads to VSDs, cardiac diastolic dysfunction and severe growth
retardation. Human ortholog, cardiomyopathy-associated 3 (CMYA3), of mXinβ is
mapped to 2q24.3. Human patients with chromosome band 2q24 deletion also exhibit
severe growth retardation and VSDs (http://www.orpha.net/data/patho/GB/uk-2q24.pdf).
The genome-wide linkage analysis of a large Kyrgyz family also reveals candidate genes
on 2q24.3-q31.1 conferring susceptibility to premature hypertension (Kalmyrzaev et al.,
2006). Further studies are warranted to characterize mXinβ’s involvement in cardiac
development, function and disease.
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Figure 3.1. Genomic structure, mRNA and protein isoforms of mXinβ. (A) The mXinβ
gene contains 9 exons (E1-E9), capable of generating 3 distinct mRNAs (mXinβ-A,
mXinβ-B and mXinβ-C), as identified by 5’ and 3’ RACE. Both mXinβ-A and mXinβ-B
mRNAs include E8 but use different poly(A) addition signals in E9. These represent the
major species, whereas the minor mXinβ-C mRNA specifically splices out the E8. (B)
Both mXinβ-A and mXinβ-B use the same stop codon (TAA) in E8 and translate into the
same mXinβ protein with 3,283 amino acid (aa) residues. The mXinβ-C uses the stop
codon (TAG) in E9 and codes for mXinβ-a protein with 3,300 aa residues. Both mXinβ
and mXinβ-a proteins contain actin-binding motifs (Xin repeats, aa#308-1,307), within
which predicted β-catenin-binding domain (β-catBD) locates. They also possess
consensus sequences for Myb DNA-binding domain (DBD), nuclear export signal (NES),
nuclear localization signal (NLS), 3 proline-rich regions (PR1, PR2, and PR3) and
ATP/GTP-binding domain (ATP/GTP-BD). (C) Western blot analysis shows that forceexpressed mXinβ protein in CHO cells has a similar mobility in SDS-PAGE gel as
endogenous mXinβ found in mouse heart extract. (D) Immunofluorescence microscopy
reveals that force-expressed mXinβ, detected by rabbit anti-mXin (U1013) and
Rhodamine-conjugated goat anti-rabbit IgG (red color), co-localizes with actin filaments
to stress fibers, labeled by fluorescein-conjugated phalloidin (green color).
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Figure 3.2. Spatial and temporal expression patterns of mXinβ in mice. (A) Western blot
analysis of total protein extracts prepared from various tissues of a postnatal day 7.5
(P7.5) mXinα-null mouse with U1013 anti-mXin antibody reveals that mXinβ is
specifically expressed in striated muscles such as tongue, heart and diaphragm. (B) The
Coomassie Blue-stained protein profile from the same total protein extracts used in (A)
shows the protein loading in each lane. (C) Western blot analysis with anti-mXin on total
protein extracts prepared from 3 individual hearts of wild type mice at P0.5, P3.5, P7.5
and P12.5 reveals a significant up-regulation of mXinβ in postnatal hearts. (D) The
Coomassie Blue-stained protein profile from the same total protein extracts used in (C)
shows the relative protein loading in each lane.
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Figure 3.3. Generation of mXinβ-null mice. (A) Targeting strategy. A restriction map of
the relevant genomic region of mXinβ is shown at the top (mXinβ locus). The targeting
vector (in the middle) contains the genomic region with a LacZ-Neor cassette to replace
portion of Exon 6 (E6)-intron 6-portion of E7. The targeted locus is shown at the bottom.
The probe used for Southern blot is located downstream of E8. (B) Southern blot analysis
of SacI-digested genomic DNAs. (C) PCR genotyping. The locations of the PCR
products for endogenous mXinβ (538bp) and targeted locus (419bp) are shown in (A).
(D) Northern blot analysis. A ~12kb mXinβ message was detected in both wild type and
heterozygous but not homozygous samples. The same membrane was hybridized with
GAPDH (glyceraldehyde 3-phosphate dehydrogenase) probe to show RNA loading. (E)
Western blot analysis on developing heart extracts from each genotype with U1013 antimXin, anti-α-actinin and anti-α-TM antibodies. A ~340 kDa mXinβ was detected in the
developing wild type and heterozygous but not homozygous extracts (the top panel).
There are no apparent changes in the expression of mXinα-a, mXinα, α-actinin, or α-TM
in the mXinβ-null hearts.
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Figure 3.4. Neither persistent truncus arteriosus (PTA) nor patent ductus arteriosus
(PDA) was detected in newborn mXinβ-null mouse heart. Ao, aorta; PA, pulmonary
artery; DA, ductus arteriosus. Bar = 1mm
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Figure 3.5. Loss of mXinβ results in severe growth retardation. (A) Body weight (BW)
comparison. * p<0.01, significant difference between mXinβ-/- and control (mXinβ+/+ or
mXinβ+/-), ANOVA. (B, C) Heart weight (HW) and HW/BW comparisons. (D, E) Liver
weight (LW) and LW/BW comparisons. The numbers of animals measured are indicated
within each bar. The means±SEM are displayed graphically. t-test for (B)~(E). N.S.,
p>0.05 no significant difference.
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Figure 3.6. Structural analyses of mXinβ+/+ and mXinβ-/- hearts. (A&B) Gross
morphology showing abnormal heart shape in P0.5 mXinβ-null mice. Bar=1mm. (C&D)
H&E-stained sections demonstrating the VSD (arrow in D) in P0.5 mXinβ-null heart. Bar
= 1 mm. (E&F) H&E-stained sections of E14.5 wild type and mXinβ-null embryos.
Bar=0.5mm. (E’&F’) higher-magnification images of the boxed areas in E&F.
Bar=50µm. (G&H) H&E-stained sections from P3.5 wild type and mXinβ-null hearts.
Bar=1mm. (G’&H’) higher-magnification images of the boxed areas in G&H.
Bar=0.1mm. (G’’&H”) higher-magnification images of the boxed areas in G’&H’,
showing mis-organized myocytes in mutant ventricle. Bar=50µm. (I&J) EM micrographs
showing no alteration in sarcomere organization of P15.5 mXinβ-null hearts. Bar=1µm.
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Figure 3.7. Doppler flow spectra recorded from the mitral valvular orifices of P12.5 wild
type and mXinβ-null mice. E-wave represents early filling velocity, whereas A-wave is
late filling (atrial contraction) velocity.
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Figure 3.8. Masson’s trichrome-stained heart sections from P11.5 wild type and mXinβnull mice demonstrating no apparent cardiac fibrosis in the mXinβ-null heart. Bar = 1mm
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Figure 3.9. Western blot analysis on total protein extracts prepared from developing
hearts of each mXinβ genotype with anti-myosin heavy chain (MHC) antibodies, anti-Ncadherin, anti-β-catenin, anti-p120-catenin and DM1B anti-β-tubulin. The MHC switch
from embryonic β-MHC to adult α-MHC in mXinβ-null hearts occurred normally. Most
of the adherens junctional components examined here was expressed normally, except
that p120-catenin may be significantly reduced in P12.5 mXinβ-null heart.
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Figure 3.10. A significant delay in switching off ssTnI in mXinβ-null hearts. (A) Protein
profiles from developing hearts of each genotype, adult soleus and transgenic mouse
hearts over-expressing cTnI-ND. (B and C) Western blot analyses with anti-TnI and anticTnT antibodies, respectively. By P7.5~13.5, the control hearts (+/+ and +/-) almost
switched off ssTnI, whereas the mXinβ-/- heart still expressed significant amounts of
ssTnI (arrows). In contrast, the cTnT isoform switch from ecTnT to acTnT in
homozygous hearts was normal. fsTnI, fast skeletal TnI.
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Figure 3.11. Increased apoptosis and decreased proliferation in developing mXinβ-null
hearts. (A&B) Representative images from P3.5 heart sections, stained with anti-active
caspase 3 for apoptotic cells (red, arrowheads), anti-cTnT for cardiomyocytes (green),
and DAPI for nuclei (blue). Bar=100µm. (A’&B’) higher-magnification images of A&B.
Bar=10µm. (C&D) Representative images from heart sections of P7.5 BrdU-labeled
mice, stained with anti-BrdU for proliferative cells (red) and counterstained with DAPI
(blue). Bar=100µm. (C’&D’) higher-magnification images of heart sections triple-stained
with anti-BrdU (red), anti-α-TM (green) and DAPI (blue). Bar=10µm. (E&F) Apoptotic
and proliferative cell populations, respectively, in developing wild type and mXinβ-null
hearts. The numbers of animals measured are indicated within each bar. The means±SEM
are displayed graphically. N.S., p>0.05 no significant difference (t-test).
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Figure 3.12. Representative heart sections from wild type (A, C) and mXinβ-null (B, D)
mice at P3.5 and P12.5. Cardiomyocytes outlined by anti-laminin (green color),
cytoplasm labeled with anti-cTnT (red color) and nuclei labeled with DAPI (blue color)
were used for comparison of cell shape difference. Irregular cell shape was readily
observed in P12.5 mXinβ-/- heart, suggesting a mis-organization in myocardium. Bar =
10 µm. (E) The cardiomyocyte width was measured by the length of the shortest axis ran
through the nuclear center of the cross-sectioned cardiomyocytes. At P3.5, there was no
apparent difference in cell width and cell shape between wild type and mXinβ-null
cardiomyocytes. In contrast, at P12.5, mXinβ-null cardiomyocytes showed a significant
reduction in cell width.
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Figure 3.13. Mis-localization of N-cadherin and mXinα as well as structural alteration in
developing intercalated disc of mXinβ-null hearts. Frozen heart sections from P16.5 mice
were immunofluorescently stained with U1040 anti-mXinβ (A&B), anti-cadherin (C&D)
and R1697 anti-mXinα (E&F). In wild type hearts, mXinβ (A), N-cadherin (C) and
mXinα (E) all localized to the mature intercalated discs. In contrast, the loss of mXinβ
(B) in mXinβ-null heart led to mis-localization of N-cadherin (D) and mXinα (F).
Bar=30µm. (G&H) EM images of P15.5 wild type and mXinβ-null cardiomyocytes.
Arrows, intercalated discs; *, T-tubules. Bar=1µm. (I&J) high magnification images of
maturing intercalated discs of P15.5 wild type and mXinβ-null cells. The closely apposite
membranes of mutant intercalated disc were less convoluted and less wavy. des,
desmosome; lig, two membranes in the process of ligation together to form intercalated
disc. Bar=0.2µm.
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Figure 3.14. The proportion of N-cadherin and connexin 43 localized to the termini of
developing cardiomyocytes of wild type and mXinβ-null mice. (A) Images of individual
cardiomyocytes immunofluorescently labeling for N-cadherin (a, b) or connexin 43 (c, d).
These images were extracted from micrographs taken from comparable regions of P15.5
wild type and mXinβ-null ventricular myocardia. The termini of cardiomyocytes were
defined by making a rectangular box at both ends of the cardiomyocytes. The width of
each box was 10% of the longitudinal axis of the cardiomyocyte as shown in (a). Both Ncadherin and connexin 43 showed higher level of terminal localization in the wild type
cardiomyocytes (a, c) than in the mXinβ-null cardiomyocytes (b, d). (B) Quantification of
terminally localized N-cadherin and connexin 43 in P15.5 and P18.5 cardiomyocytes. In
both wild type and mXinβ-null hearts, the assembly of Cx43 to the myocyte termini was
well behind the assembly of N-cadherin at both time points (p<0.02). The percentages of
terminally localized N-cadherin and connexin 43 in the wild type cardiomyocytes at both
time points were significantly higher than that in the mXinβ-null counterparts. In addition,
the terminal localization of N-cadherin and connexin 43 increased significantly from
P15.5 to P18.5 in the wild type cardiomyocytes, whereas the terminal localization of both
proteins in the mXinβ-null cardiomyocytes remain unchanged during the same
developmental stage. The number of cardiomyocytes measured was indicated in each bar.
* p<0.05. N.S., no significant difference.
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Figure 3.15. Increased Stat3 activity and decreased Rac1, IGF-1R, Akt and Erk1/2
activities in mXinβ-null hearts. (A) The relative GTP-bound Rac1 but not GTP-bound
RhoA was significantly reduced in P7.5 mXinβ-null hearts. N.S., p>0.05 no significant
difference (ANOVA). The numbers of animals measured are indicated within each bar.
(B) Key signaling pathways involved in cell proliferation, growth and survival in the
heart. (C) Western blot analyses with phospho-specific antibodies against key signaling
molecules. Up-regulation of active Stat3 (p-Stat3) (^) can be detected in some of mXinβnull hearts as early as P0.5. However, this Stat3 activation is not parallel to the activation
of Jak2 (p-Jak2). Total Akt and Grb2 protein levels remain unchanged; however, the
activation/phosphorylation of Akt, GSK3β, Erk1/2 and IGF-1R were significantly
reduced beginning from P7.5 in mXinβ-null hearts (*).
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Figure 3.16. No mis-localization of mXinβ in mXinα-null mouse heart.
Immunofluorescence microscopy was performed on frozen heart section from adult
mXinα-null mice with affinity-purified rabbit U1040 anti-mXinβ (red color) and mouse
monoclonal anti-β-catenin (green color). The nuclei were labeled with DAPI (blue
color). Bar = 10µm
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Figure 3.17. Proposed roles of mXinβ in postnatal heart growth. In the wild type heart, Ncadherin and its associated proteins mediate bi-directional signaling and cross-talks,
because these proteins are shown to interact with many signaling molecules such as
receptor tyrosine kinases (e.g., IGF-1R), non-receptor tyrosine kinases (e.g., c-Src, Jak),
tyrosine phosphatases, phosphatidylinositol-3 kinase (PI3K) and adaptors (Braga and
Yap, 2005; McLachlan and Yap, 2007; Pece et al., 1999; Wheelock and Johnson, 2003;
Xu et al., 1997). The pleiotropic effects caused by deletion of mXinβ suggest that mXinβ
is a pivotal factor for both N-cadherin-mediated bi-directional and cross-talk signalings.
Similar to mXinα (Choi et al., 2007), the mXinβ containing several conserved binding
domains may also interact with β-catenin, p120-catenin and actin filaments. Together,
mXinα and mXinβ may play important role in the Rac1 activation through c-Src-PI3K
and Vav2 (a guanine nucleotide exchange factor capable of binding to p120-catenin),
similar to the signaling found in epithelial cells (Fukuyama et al., 2006; Noren et al.,
2000; Noren et al., 2001). Postnatal heart growth requires an intricate regulation of Rac1
activity (Sussman et al., 2000), and the Rac1 activation is essential for rearranging actin
cytoskeleton to align cells in response to mechanical stretch (Yamane et al., 2007) and for
modulating mitogen-activated protein kinase activity and myocardial oxidative stress
(cross-talk signaling) in response to various hypertrophic stimuli (Clerk et al., 2001;
Satoh et al., 2006). In the mXinβ-null heart, the loss of mXinβ impairs the engagement
and clustering of N-cadherin, down-regulates the Rac1 activity, and subsequently mislocalizes mXinα. These impairments would in turn dys-regulate hormone-, cytokine-, and
growth factor-mediated signalings for postnatal heart growth. Although the mechanism
remains to be determined, mXinβ-null hearts exhibit a persistent activation of Stat3 and a
down-regulation of IGF-1R activity. Furthermore, the up-regulation of Stat3 activity in
mutant hearts appears to precede the reductions in the activities of growth-related
signaling molecules. Since the stat3 activity is auto-regulated by many positive (such as
Jak, c-Src, IGF-1(Honsho et al., 2009)) and negative (such as suppressor of cytokine
signaling protein 3, SOCS3 (Kurdi and Booz, 2007), cytoplasmic tyrosine phosphatases)
regulators (Boengler et al., 2008; Levy and Darnell, 2002), it should be worthy to
determine which of these regulators, including the ATP/GTP-binding domain-containing
mXinβ, are responsible for the up-regulation of Stat3 activity. The defect in the clustering
of N-cadherin in mXinβ-null hearts may also impair the IGF-1R organization during
postnatal heart growth, leading to the reduced activities of IGF-1R, Akt and Erk-1/2, and
the severely retarded growth.
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Table 3.1. Genotypes of progenies of mXinβ+/- intercrosses
Age
P0.5
P3.5
P7.5
P12.5
P17.5
Number of mice observed(number of mice
expected)
mXinβ+/+
mXinβ+/mXinβ-/46 (45.5)
101 (91)
35 (45.5)
57 (38)
72 (76)
23 (38)
50 (38.25)
79 (76.5)
24 (38.25)
37 (29.25)
68(58.5)
12 (29.25)
12 (10)
28 (20)
0 (10)
Total
number
182
152
153
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40
p value
0.17
<0.01
0.01
<0.01
0.02
Age: at which genotype was determined.
The number of mice observed is indicated for each genotype and the number of mice
expected from Mendelian frequency is shown in parenthesis.
p value: determined from Chi square test.
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Table 3.2. Echocardiographic analysis of control (mXinβ+/+ & mXinβ+/-) and mXinβnull mice at P3.5 and P12.5
Parameters
Heart rate (bpm)
IVSd (mm)
IVSs (mm)
LVPWd (mm)
LVPWs (mm)
LVIDd (mm)
LVIDs (mm)
LVVd (µl)
LVVs (µl)
EF (%)
FS (%)
Mitral valve E/A
ratio
P3.5
Control (n=7)
mXinβ-/- (n=5)
458±48
0.44±0.08
0.73±0.09
0.48±0.13
0.66±0.10
1.63±0.10
0.83±0.11
7.65±1.18
1.36±0.57
83.22±4.68
49.34±5.65
0.83±0.03
473±59
0.47±0.03
0.77±0.14
0.42±0.03
0.74±0.15
1.45±0.21
0.60±0.13*
5.76±2.01
0.60±0.26*
88.37±6.15
56.69±11.19
0.73±0.06*
P12.5
Control (n=4)
mXinβ-/- (n=5)
472±47
0.57±0.10
0.78±0.14
0.62±0.16
0.89±0.16
2.53±0.16
1.61±0.21
23.07±3.55
7.60±2.24
67.35±7.47
36.07±5.78
1.64±0.10
444±58
0.46±0.06
0.77±0.10
0.56±0.18
0.76±0.14
2.10±0.13*
1.27±0.23
14.47±2.28*
4.21±2.06*
72.37±9.54
40.81±8.74
1.11±0.19*
Two dimensional images were recorded in parasternal long- and short-axis projections
with guided M-mode recordings at the midventricular level in both views. Left ventricle
(LV) chamber size and wall thickness are measured in at least three beats from each
projection and averaged.
bpm: beats per minutes
IVSd and IVSs: interventricular septum thickness at diastole and systole, respectively.
LVPWd and LVPWs: LV posterior wall thickness at diastole and systole, respectively.
LVIDd and LVIDs: LV internal dimension at diastole and systole, respectively.
LVVd and LVVs: LV volume at diastole and systole, respectively.
EF: ejection fraction
FS: fraction shortening
E/A: mitral valve E-wave (early filling) to A-wave (atrial contraction/late filling) ratio
* p ≤ 0.05 significant difference between mXinβ-/- and control mice (Student’s t-test)
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Table 3.3. Assessment of ventricular myoarchitecture
mXin+/+
mXin-null
(n=5)
(n=4)
240.4±15.2
188.5±7.5
0.026*
Left ventricle trabecular region
41.1±4.9
51.3±9.1
0.332
Right ventricle compact region
168.3±16.8
120.0±18.6
0.096
67.3±6.1
114.0±23.7
0.070
Area/length (m2/m)
Left ventricle compact region
Right ventricle trabecular region
p value
Compact and trabecular regions per unit myocardium within lateral free walls of left and
right ventricles of newborn wild type and mXin-null mice were determined from H&E
stained sections according to the previously described method.(Ishiwata et al., 2003) This
analysis used in developing mouse embryos has previously revealed that the
developmental changes in ventricular myoarchitecture correlate very well with the
changes in ventricular diastolic function. In general, peak E-wave velocity (active
relaxation) is exponentially correlated with the area of compact region per 1m
myocardium, while peak A-wave velocity (passive compliance/atrial contraction) is
correlated with the area of trabecular region per 1 m myocardium. A significant
decrease in the left ventricle compact region detected in P0.5 mXin-null hearts is
consistent with a smaller E-wave velocity and thus diastolic dysfunction.
* significant difference between mXin+/+ and mXin-/- hearts (Student’s t-test)
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CHAPTER IV
mXINβ IS ESSENTIAL FOR THE POSTNATAL MATURATION OF
THE INTERCALATED DISCS
Preface
The data presented in this chapter will be included in a manuscript by Qinchuan
Wang, Jenny Li-Chun Lin and Jim Jung-Ching Lin (in preparation). In this study, I
provide multiple lines of evidence to support that mXinβ is essential for postnatal
development of the intercalated discs (ICDs). I used quantitative Western blot to examine
the temporal expression profiles of mXinα, mXinβ as well as the core adherens junction
protein N-cadherin. These experiments revealed that the temporal expression profile of
mXinβ correlates very well with the process of ICD maturation and the onset of ICD
defects in mXinβ-null hearts. Furthermore, using immunofluorescence staining and
subcellular fractionation, I showed that mXinβ is specifically associated with the
maturing ICDs, suggesting mXinβ functions initially and locally to promote ICD
maturation. This study also provides evidence to show that mXinβ is essential for the
distribution of intercellular junction components at the cellular level, but it is not require
for their associations among N-cadherin, desmoplakin and connexin 43, suggesting
multiple mechanisms are responsible for the establishment of the intricate ICDs.
Abstract
Intercalated discs (ICD) are cardiac-specific structures responsible for
mechanical, electrical and chemical communication between cardiomyocytes and are
implicated in signal transduction. Defects of ICD components cause a number of human
cardiac diseases, and changes of ICDs are associated with cardiomyopathy, arrhythmias,
and heart failure. ICDs are formed during postnatal development through a profound
redistribution of the intercellular junctions and recruitment and assembly of more than
200 proteins at the termini of cardiomyocytes. The molecular mechanism of this process
157
is unclear. The mouse orthologs (mXinα and mXinβ) of human cardiomyopathyassociated genes (CMYA1 and CMYA3, respectively) encode proteins localized to ICDs.
Previously, we showed that ablation of mXinα results in adult late-onset cardiomyopathy
with conduction defects and up-regulation of mXinβ, and ICD structural defects are
found in adult but not juvenile mXinα-null hearts. On the other hand, loss of mXinβ leads
to ICD defects at postnatal day 16.5, a developmental stage when the heart is forming
ICDs, suggesting mXinβ is required for ICD maturation. In this study, with quantitative
Western blot, we showed that mXinβ but not mXinα is uniquely upregulated during the
redistribution of intercellular junction from the lateral membrane of cardiomyocytes to
the cells’ termini. Loss of mXinβ leads to failure of restricting the intercellular junctions
to the termini of the cells, and the onset of such defect correlates with the peak expression
of mXinβ. Immunofluorescence staining and subcellular fractionation showed that
mXinβ preferentially associates with the maturing ICDs, further suggesting that mXinβ
functions locally to promote ICD maturation. In contrast, the spatiotemporal expression
profile of mXinα, and the lack of more severe ICD defects in mXinα-/-:mXinβ-/- double
mutant hearts than in mXinβ-/- hearts suggest that mXinα is not essential for the postnatal
maturation of ICDs.
Introduction
The integration of the contraction and relaxation of billions of individual
cardiomyocytes is essential for the heart to function. To carry out such integration,
individual cardiomyocytes must be excited at the right moment so that they have
coordinated contractions in each heartbeat, which requires electrical coupling between
cardiomyocytes. The contractile forces generated by individual cardiomyocytes in turn
must be transmitted to the correct neighbors so that the tiny forces from each
cardiomyocytes are added up for the heart to perform mechanical work, which requires
mechanical coupling between cardiomyocytes. These essential electrical and mechanical
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couplings are carried out by a cardiac specific structure, the intercalated discs (ICD). The
functions of ICDs have been attributed to three types of intercellular junctions (Forbes
and Sperelakis, 1985): the gap junctions that are made of connexin permit ions to flow
between cardiomyocytes for electrical coupling; the adherens junctions that are organized
by N-cadherin confers continuity for the myofibrils between cardiomyocytes and are
central for transmitting contractile forces; and the desmosomes that are organized by the
desmosomal cadherins couple the sarcolemma to the intermediate filaments to maintain
the mechanical integrity of the cardiomyocytes. In addition to these functions classically
assigned to the intercellular junctions of the ICDs, it is now increasingly realized that
ICDs are specialized membrane domains of the cardiomyocytes with more than 200
proteins (Estigoy et al., 2009) and also function in chemical and mechanical signaling as
well as ion transportation (Noorman et al., 2009).
Given the important roles of ICDs, it is not surprising that mutation of genes
encoding ICD components can cause severe heart diseases, such as the arrhythmogenic
right ventricular cardiomyopathy (Delmar and McKenna, 2010). Conversely, various
heart diseases not directly related to mutations of ICD components lead to alterations in
the ICDs, and such alterations are likely an important facet of the pathology of these
diseases (Barker et al., 2002; Noorman et al., 2009; Wang and Gerdes, 1999). The
important roles of the ICDs in the health and disease of the heart are further supported by
the severe cardiac defects of a number of animal models (Ferreira-Cornwell et al., 2002;
Kostetskii et al., 2005; Li et al., 2006; Li and Radice, 2010; Wang et al., 2010). An
important theme of these cardiac diseases and defects, either originated from mutations of
ICD components or from non-ICD related reasons is that the molecular integrity of the
ICDs are disrupted, which is manifested as altered morphology, abnormal expression
levels and localizations of protein components as well as changed protein-protein
interaction profiles of the ICDs. This fact indicates that the precise structure and
organization of the ICDs are vital for them to carry out their specific functions.
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The components of the complex ICDs are assembled mainly during postnatal
development as ICDs mature. Several studies have shown that the maturation of ICDs is
characterized by drastic reorganization of the distribution of intercellular junctions (Angst
et al., 1997; Hirschy et al., 2006; Peters et al., 1994). In the embryos, immunostaining
showed that N-cadherin and associated proteins such as β-catenin and mXinα are
localized to almost the entire surface of cardiomyocytes in a rather diffused pattern (Sinn
et al., 2002). Gap junction and desmosomal proteins are also distributed on the entire
surface of cardiomyocytes where cell-cell contacts exist but show more spotted
localization than the components of adherens junctions (Coppen et al., 2003; Pieperhoff
and Franke, 2007). During postnatal development, the intercellular junctions undergo
reorganization by which all three types of intercellular junctions are eventually localized
to the ends of cardiomyocytes. Partially overlapping with the reorganization of the
geometry of the junctions at the cellular level is the intermixing of the adherens junctions
with the desmosomes at the molecular level to form a newly identified structure, area
composita (Pieperhoff and Franke, 2007).The molecular mechanism for the postnatal
maturation of ICDs is largely unknown. However, since all the classic components of the
intercellular junctions are already expressed in the embryonic heart, the postnatal
reorganization of the intercellular junctions for the maturation of ICDs must be dictated
by additional factors that are expressed/activated during the postnatal life.
One such factor might be the intercalated protein mXinβ, which is a member of
the Xin repeat-containing family of proteins that are specifically localized to the ICDs in
the adult cardiomyocytes (Lin et al., 2005). In the mouse, the Xin repeat-containing gene
family has two members, the mXinα and mXinβ, which encodes the mXinα alternatively
splicing variants (mXinα and mXinα-a) and mXinβ alternatively splicing variants
(mXinβ and mXinβ-a) respectively (Gustafson-Wagner et al., 2007; Wang et al., 2010).
The Xin repeats are conserved protein motifs that interact with actin filaments (Choi et
al., 2007; Pacholsky et al., 2004). Within its Xin repeat region, the mXinα variants have a
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conserved β-catenin-interacting domain (Choi et al., 2007). This β-catenin-interacting
domain and its counterpart in mXinβ may be responsible for recruiting both the mXin
proteins to the adherens junctions, where the mXin proteins may directly couple the Ncadherin-catenin complex to the underlining actin cytoskeleton (Grosskurth et al., 2008).
Evidence from our previous study indicates that mXinβ may play important roles
in the postnatal maturation of ICDs. We observed an up-regulation of mXinβ protein in
the heart from P0.5 to P13.5 (postnatal day 0.5 and 13.5 respectively); we also found that
in the mXinβ-/- hearts at P16.5, the intercellular junctions are punctate and mature ICDlike structures are sparse (Wang et al., 2010). However, several gaps in our knowledge
prevent us from drawing a firm conclusion about mXinβ’s roles in ICD maturation. First,
although previous studies showed that ICDs are formed postnatally, descriptions of the
reorganization of intercellular junctions between P0.5 and P16.5 are not detailed enough
for us to correlate this process with the expression profile of mXinβ. Second, the
spatiotemporal expression profile was not fully characterized for mXinβ: we don’t know
the expression profile of mXinβ after P13.5 and whether mXinβ is localized to the
adherens junctions throughout their reorganization. Third, the nature of the defects of
ICDs in mXinβ-/- hearts was not addressed in our previous study: we don’t know whether
the ICDs are not formed at all in the mXinβ-/- hearts or are formed but then fail to be
maintained. In this study, we asked what specific roles mXinβ plays in the maturation of
ICDs and answered this question by investigating the quantitative expression profiles of
mXinβ and the core protein of adherens junctions, N-cadherin; we also provided a
detailed description of the time course of reorganization of the intercellular junctions
during ICD maturation in wild-type and mXinβ-/- hearts. Our results suggest a direct
involvement of mXinβ in the postnatal reorganization of intercellular junctions.
In addition to studying mXinβ’s roles in the maturation of ICDs, we examined
mXinα variants’ roles in this process because mXinα variants share many conserved
regions with mXinβ but seems incapable of compensating for the loss of mXinβ for the
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maturation of ICDs (Wang et al., 2010). This is in contrast with the compensatory roles
of mXinβ for the loss of mXinα we demonstrated previously (Gustafson-Wagner et al.,
2007). We asked whether the inability of mXinα variants to compensate for the loss of
mXinβ is due to insufficient expression during ICD maturation or lack of essential protein
function required for this process.
Materials and Methods
Animals
All animal procedures were approved and performed in accordance with
institutional guidelines.
Antibodies
Primary antibodies: rabbit polyclonal antibodies against both mXinα and mXinβ
(U1013), mXinα specifically (R1697), and mXinβ specifically (U1040) were generated in
our lab and reported previously (Sinn et al., 2002; Wang et al., 2010). Other antibodies
were purchased from commercial sources: mouse anti-N-cadherin (3B9, Invitrogen, used
for immunofluorescence staining), rat anti-N-cadherin (MNCD2, Developmental Studies
Hybridoma Bank, for Western blot), rabbit anti-connexin 43 (Zymed Laboratories Inc.),
rabbit anti-desmoplakin (AHP320, Serotec), mouse anti-β-catenin (CAT-5H10, Zymed
Laboratories Inc.), mouse anti-p120ctn (15D2, Zymed Laboratories Inc.), and mouse antiGAPDH (6C5, RDI Research Diagnostics, Inc.).
Secondary antibodies: Dylight 488 conjugated goat anti-mouse (Thermo
Scientific) and Cy5 conjugated goat anti-rabbit (Chemicon International Inc.) were used
for immunostaining. IRDye 800CW conjugated goat anti-rat (ODYSSEY), IRDye 800
conjugated goat anti-mouse (Rockland) and IRDye 800 conjugated Goat anti-rabbit
(Rockland) were used for Western blot.
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Quantitative Western blot
To generate recombinant protein as standard for quantification of mXinβ, we
subcloned a cDNA fragment of mXinβ encoding the entire region recognized by anti-Xin
antibody U1013 (aa#1 – 1293) from the previously described full-length cDNA into
pGEX4-T2 vector (GE Healthcare Life Science). The GST-fused mXinβ fragment
predicted to have molecular weight of 177.1 kD was named GST-mXinβ5’. The GSTmXinβ5’ was expressed in BL21(DE3)pLysS bacteria and affinity purified with
Glutathione Sepharose 4B beads (Amersham). To determine the concentration of the
intact fragment (177.1 kD) of purified GST-mXinβ5’, we subjected the purified protein to
SDS-polyacrylamide gel electrophoresis (SDS-PAGE) alongside with serially diluted
Bovine Serum Albumin protein standards (BSA, Sigma). The gel was stained by
Coomassie Brilliant Blue solution for 2 hours and then de-stained and imaged by EC3
imaging system (Ultra-Violet Products, Ltd). The intensities of the protein bands were
quantified with NIH ImageJ (http://rsbweb.nih.gov/ij/) and the concentration of the GSTmXinβ5’ was determined against the intensity-concentration standard curves established
with BSA protein standards.
For quantification of mXinβ, heart samples and GST-mXinβ5’ standards were
loaded into 6% SDS-polyacrylamide gels. For each postnatal stage, samples (each
contains 0.125 mg of tissue) from three different hearts were loaded into each gel, and
each heart sample was loaded into two lanes as duplicates. In order to generate standard
curves, 12.5 ng, 6.25 ng, 3.13 ng and 1.56 ng of GST-mXinβ5’ were loaded into separate
lanes in each gel alongside the heart samples. Following electrophoresis, the proteins
were transferred overnight at 20 volts to nitrocellulose membranes (Millipore). Then,
Western blot experiments were carried out following the Western Blot Analysis protocol
from Li-Cor Biosciences, using U1013 as the primary antibody. The blots were imaged
by an Odyssey Imager and the images were analyzed by ImageJ. One standard curve was
generated for each blot by plotting the amount of GST-mXinβ5’ in each lane against the
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signal intensities of the corresponding protein bands in Western blot. Based on the
standard curve, the amount of endogenous mXinβ in each heart sample was calculated.
To quantify mXinα variants, we used purified GST-mXinα5’(86.7 kD) as
standard. All procedures were the same as the above description for the quantification of
mXinβ except that each of the samples loaded for Western blot contains contents from
0.0125 mg of heart tissue. The amount of GST-mXinα5’ used for standard were 1.25 ng,
0.625 ng, 0.313 ng and 0.156 ng. U1013 was used for Western blot detection.
To quantify N-cadherin, a purified recombinant human N-cadherin extracellular
region (Sino Biological Inc) that was expressed in Chinese Hamster Ovary cells as
secreted protein was used to generate standard curves. This commercial product contains
both the pro- and mature form of the N-cadherin’s extracellular region and runs as two
bands (90 kD and 75 kD) in SDS-PAGE under reducing condition. We quantified the
mature 75 kD form with BSA standard as described above. For quantitative Western blot,
the loading of the heart samples was identical to the ones in the mXinα quantification
experiments. To generate standard curve, 5.00 ng, 2.50 ng, 1.25 ng and 0.625 ng of the
N-cadherin recombinant protein were used. The primary antibody used for the Western
blot was the Rat-anti N-cadherin monoclonal antibody (MNCD2, Developmental Studies
Hybridoma Bank).
Immunostaining
Immunostaining were carried out on 4-µm frozen sections. The sections were
fixed in 3.7% formaldehyde in phosphate buffered saline (PBS) for 10 minutes at room
temperature, rinsed with PBS and permeabilized with cold acetone (-20 ºC) for 5 minutes.
After blocking the sections with the blocking reagent from the Vector Laboratories’s
Mouse on Mouse kit for 1 hour at room temperature and followed by blocking with 5%
normal goat serum for 30 minutes, the sections were incubated with primary antibodies
diluted with the Pierce Immunostain Enhancer (Thermo Scientific) for 30 minutes at 37
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ºC. The sections were then washed with PBS and incubated with secondary antibodies
diluted in the Pierce Immunostain Enhancer for 10 minutes at 37 ºC. After wash, the
sections were sealed in antifading reagent (Gelvatol) and covered by coverslips. Sections
were imaged with Leica TCS SPE confocal microscope with an ACS APO 40x N.A. 1.15
oil objective.
Quantification of confocal images
Quantification was carried out with ImageJ. Briefly, confocal images of
longitudinally sectioned cardiomyocytes were first randomly selected. The total
integrated immunofluorescence signals (pixel number X intensity) were measured by
ImageJ after using its default threshold function. Then the ICD/ICD-like signals (defined
as signal clusters that have one dimension as wide as the width of the cardiomyocyte’s
terminus where the signals reside) were masked by black pixels in the images and the
integrated immunofluorescence signals (non-ICD) were measured again. The ICD-signal
was then calculated by subtracting the non-ICD signals from the total signals.
For quantification and statistical test of the ICD/total signal ratio, we used at least
5 confocal images of each heart and for each stage, at least two control hearts and two
mutant hearts were used. The ICD/total signal ratio of each image was treated as a sample
when carrying out Student’s t-test.
To examine the association between connexin 43 and N-cadherin signals, we used
the find maxima function of ImageJ to determine the x- and y- coordinates of each signal
spot in confocal images and output the coordinates into Microsoft Excel. To find the
closest N-cadherin spot for a specific connexin 43 spot, the minimal distance from all Ncadherin spots in the same images were determined for the specific connexin 43 spot with
a custom Excel macro, RED/GREEN DOT PROCESSOR (Appendix A).
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Subcellular fractionation
Subcellular fractionation experiments were carried out following a modified
protocol that was used in isolating ICDs from large quantity of adult heart tissues (Colaco
and Evans, 1981). Briefly, one or two hearts were homogenized in ice-cold
homogenization buffer (10 mM Imidazole buffer, pH7.0, containing complete protease
inhibitor cocktail from Roche) with a Multigen 7 homogenizer for 2.0 min at full-speed
and followed by Dounce treatment with pestle B (loose fitting) for 30 times. The
homogenate was layered on top of a sucrose step gradient made with the homogenization
buffer in the Beckman Ultra-Clear Tubes (14 x 89 mm). The steps of the gradient from
the bottom to the top of the tubes were: 51.5% (w/w), 46.5%, 41.5%, 36.0%, 30.0% and
23.0%. After centrifugation for 20 hours at 28,000 RPM with an SW41 rotor, 20 fractions
were collected from the bottom of the tube by puncturing the tube with a syringe needle.
The fractions were analyzed by Western blot following a standard protocol.
Results
The adherens junction proteins, mXinβ, mXinα and Ncadherin have unique temporal expression profiles during
postnatal development
(i) mXinβ has a dynamically regulated three-phase
expression profile characterized by a unique and drastic upregulation between P0.5 and P13.5
To determine the roles of mXinβ in the maturation of ICDs, we first examined
mXinβ’s temporal expression profile in the mouse heart during postnatal stages from the
initiation to the completion of the maturation process of ICDs (P0.5 to P60.5). The
mXinβ’s expression profile was established by quantitative Western blot experiments,
which allowed us to determine the molar amount of mXinβ expressed in the heart, and
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thus facilitated comparisons of the expression of mXinβ among different developmental
stages and to that of N-cadherin and mXinα. Using this approach, we measured the
amount of mXinβ expressed in the heart at multiple time points from P0.5 to P60.5, with
three hearts for each time point (Figure 4.1 A and B). The data (Figure 4.1 A) showed
that during this period, mXinβ’s expression in the heart fluctuates between 0.185 ± 0.025
x 10-12 mol (all data presented as mean ± SE) per mg heart at lowest expression level
(P20.5) and 0.592 ± 0.078 x 10-12 mol per mg heart at highest expression level (P13.5).
Since mXinβ is expressed specifically in the cardiomyocytes, which make up the
majority of the volume of the heart (Legato, 1979), the concentration of mXinβ in the
cardiomyocytes is estimated to be about 0.2 µM to 0.6 µM. However, because mXinβ is
not uniformly localized in the cardiomyocytes, the local concentration of mXinβ at the
ICDs could be much higher. Interestingly, this estimated concentration of mXinβ in the
cardiomyocytes is on par with the estimated cytosolic concentration (0.6 µM) of another
adherens junction protein, α-catenin in the epithelial MDCK cells (Drees et al., 2005). By
plotting the expression level of mXinβ against the age of the mice (Figure 1 A and B),
our data revealed a three-phase expression profile of mXinβ between P0.5 and P60.5: a
rapid up-regulation between P0.5 and P13.5, followed by a sharp down-regulation
between P13.5 and P20.5, and then an up-regulation from P20.5 to P60.5. The relative
changes of mXinβ expressed in each mg of heart are +172.6% (P0.5 to P13.5), -68.8%
(P13.5 to P20.5) and +258% (from P20.5 to P60.5) respectively. Corrected with the heart
weights, corresponding folds of change of mXinβ in each heart in the above stages are
12.1, -0.65 and 3.14 respectively. Thus, between P0.5 and P60.5, the expression of
mXinβ is dynamically regulated. The drastic up-regulation of mXinβ both in
concentration and total amount between P0.5 and P13.5 suggests that mXinβ may play
important roles during this period.
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(ii) mXinα variants have a four-phase expression profile
characterized by relatively constant concentrations between
P0.5 and P13.5.
Since the young mXinα-null animals have normal ICDs, we asked whether the
mXinα variants would have a differently expression profile compared to mXinβ during
the period of ICD maturation. We carried out quantitative Western blot experiments to
determine the expression profiles of both mXinα variants with the same heart samples
used for the mXinβ experiments. The data showed that each mXinα variant is expressed
at about 5 fold the level of mXinβ at P0.5, and the difference between mXinα variants
and mXinβ becomes smaller as the heart matures (Figure 4.1 C and D). The mXinα’s
amounts per mg of heart range from 0.603 ± 0.057 x 10-12 mol at P60.5 to 1.331 ± 0.146
x 10-12 mol at P7.5; the mXinα-a amounts per mg of heart range from 0.383 ± 0.010 x 1012
mol at P20.5 to 1.137 ± 0.027 x 10-12 mol at P3.5. The concentrations of mXinα
variants in the cardiomyocytes are estimated to be between 0.4 µM to 1.3 µM. Plotting
the amount of mXinα variants per mg of heart against age shows that the expression of
mXinα variants between P0.5 and P60.5 has four phases (Figure 4.1 C), which is distinct
from the three-phase expression of mXinβ (Figure 4.1 A). Specifically, from P0.5 to
P13.5, mXinα variants are expressed at relatively constant levels per mg of heart (Figure
4.1 C), with only 7.8% up-regulation and 16.5% down-regulation per mg of heart tissue
for the mXinα and mXinα-a respectively, which are much smaller changes than the
172.6% increase of mXinβ during the same period. From P13.5 to P20.5, the amount of
mXinα and mXinα-a expressed per mg of heart tissue decrease by 50.0% and 58.7%
respectively, comparable with the 68.8% down-regulation of mXinβ during the same
period. After P20.5, the mXinα variants initially up-regulate and reached a peak at p30.5
but then down-regulate again, in contrast with the continuous up-regulation of mXinβ
from P20.5 to P60.5 (Figure 4.1 C). The profiles of the total amounts of mXinα variants
in each heart are accordingly very different from that of mXinβ (Figure 4.1 D).
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Interestingly, although mXinα and mXinα-a are expressed at almost the same levels at
P0.5 (1.14 ± 0.006 x 10-12 mol per mg heart and 1.11 ± 0.029 x 10-12 mol per mg heart
respectively), their expression diverge at P3.5. mXinα-a is expressed at lower levels than
mXinα from P3.5 onward, suggesting the mXinα variants may play different roles in
prenatal and postnatal life. In summary, mXinα variants have distinct profiles of
expression compared to mXinβ. In particular, concentrations of mXinα variants per mg
heart remain relatively constant between P0.5 and P13.5, similar to the expression profile
of N-cadherin (Figure 4.1 E). These suggest that mXinα variants may play certain
constitutive roles during this period but they are unlikely to be directly involved in
initiating specific developmental changes, such as ICD maturation, that start within this
period.
(iii) Total amount of N-cadherin exponentially increases
and reaches plateau at P15.5
The dynamic expression profiles of mXinβ and mXinα variants promoted us to
ask whether their expression profiles reflect the change of the expression of the core
adherens junction protein, N-cadherin. Thus, we quantified the expression profile of Ncadherin with the above heart samples by quantitative Western blot experiments. Plotting
the expression of N-cadherin per mg of heart tissue against age shows that N-cadherin
level follows a slowly reducing trend from P0.5 to P60.5 with minor fluctuations during
the process (Figure 4.1 E). From P0.5 to P13.5, N-cadherin expression per mg heart
reduces slightly by 8.9% (from 2.486 ± 0.092 x 10-12 mol to 2.264 ± 0.085 x 10-12 mol
per mg of heart). By P60.5, N-cadherin is expressed at 1.501 ± 0.137 x 10-12 mol per mg
of heart, a 39.6% reduction from P0.5. When the total N-cadherin expressed in each heart
is plotted against age, it is clear that expression of N-cadherin has two phases: it rapidly
increases from P0.5 to P15.5, and then remains almost unchanged from P15.5 to P60.5
(Figure 4.1 F). The expression profile of N-cadherin seems to reflect the postnatal growth
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of the heart: plotting the weights of the hearts used for the quantitative Western blot
against age shows that postnatal growth of the heart also has two phases, a rapid growth
phase from P0.5 to P15.5 followed by a slow growth phase from P15.5 to P60.5 (not
shown). The transition of the growth phases of the heart at P15.5 coincides with the time
point when N-cadherin expressed in each heart reaches the plateau. It is remarkable that
after P15.5, N-cadherin per heart level is very stable despite the slow, albeit continuous
growth of the heart. Thus, the data show that the expression profiles of mXinβ and
mXinα variants do not simply follow that of N-cadherin; the unique rapid up-regulation
of mXinβ between P0.5 and P13.5 likely reflects important roles mXinβ plays during this
period.
(iv) Total amount of mXin proteins is similar to that of Ncadherin except at P20.5
We also directly compared the total amount of mXin proteins with that of Ncadherin (Figure 4.2) because all the mXin proteins might associate with N-cadherin
through β-catenin, thus a quantitative relationship might exist. Interestingly, between
P0.5 and P60.5, the total amount of mXin proteins is similar to that of N-cadherin in
majority of the time points we studied. The correlation between the total levels of the
mXin proteins and N-cadherin suggests N-cadherin may influence the expression of
mXin and/or mXin proteins may influence the level of N-cadherin. The comparison also
revealed that mXinβ is only expressed at a small fraction of N-cadherin. At P0.5, mXinβ
is only expressed at 8.71% of the level of N-cadherin; this number increased to 26.6% at
P13.5, when mXinβ expression reaches a peak. Such quantitative relationship indicates
that even if a majority of mXinβ associates with N-cadherin/β-catenin complexes, only a
minor population of N-cadherin/β-catenin complexes could have mXinβ as their partners.
Because mXinβ is required for the development of normal ICDs by P16.5, we asked if
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mXinβ is specifically associated the populations of N-cadherin that are being
incorporated into the maturing ICDs.
mXinβ but not mXinα variants preferentially associates
with a subpopulation of N-cadherin at the maturing ICDs
To determine if mXinβ is specifically associated with the population of Ncadherin that is incorporating into the maturing ICDs, we did double-label
immunofluorescence staining of mXinβ and N-cadherin on frozen sections of postnatal
hearts. Representative confocal images of P7.5 and P24.5 sections are shown (Figure
4.3). We found that at P7.5, N-cadherin is distributed extensively on the surface of
cardiomyocytes (Figure 4.3 A and C). On the lateral surface of the many cardiomyocytes,
N-cadherin staining is characterized by almost continuous signal interspaced with
strongly stained puncta. Larger N-cadherin clusters can be found on the termini of the
cardiomyocytes, where the ICDs are being formed (Figure 4.3 A, arrow). Interestingly, at
this stage, mXinβ is sparse and associates with the bright N-cadherin puncta that are
found mainly at the longitudinal termini of the cardiomyocytes (Figure 4.3 B and C). A
similar phenomenon was found in both P3.5 and P13.5 hearts (data not shown). Thus,
during ICD maturation only a subpopulation of N-cadherin-containing complexes are
associated with detectable level of mXinβ, and importantly, the mXinβ-N-cadherin colocalization was primarily found at the maturing ICDs located at termini of
cardiomyocytes. At P24.5, the laterally localized N-cadherin largely disappears and
almost all N-cadherin signals are found to be highly co-localized with mXinβ at the
termini of the cardiomyocytes (Figure 4.3 D, E and F). Thus, during ICD maturation,
mXinβ is preferentially associated with a subpopulation of N-cadherin at the maturing
ICDs located at the termini of cardiomyocytes. On the other hand, double labeling of Ncadherin and mXinα/mXinα-a shows that at P7.5 (Figure 4.4 A, B and C) a majority of
the N-cadherin signal overlaps with the mXinα/mXinα-a signal; only a few lateral surface
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areas showed faintly stained N-cadherin but no mXinα/mXinα-a. Importantly,
mXinα/mXinα-a seems to have no preference for the maturing ICDs at the termini of the
cardiomyocytes. At P24.5, N-cadherin is highly co-localized with mXinα/mXinα-a in the
ICDs (Figure 4.4 D, E and F). Thus, the results show that only mXinβ, but not
mXinα/mXinα-a preferentially associates with the subpopulation of N-cadherin at the
termini of cardiomyocytes where the ICDs are being formed, further supporting mXinβ’s
specific roles in the maturation of ICDs.
mXinβ preferentially associates with a subcellular fraction
containing the maturing ICDs
We provide an additional line of evidence to support the preferential association
of mXinβ with the subpopulation of N-cadherin incorporating into the mature ICDs by
subcellular fractionation of developing postnatal hearts. Total homogenates of hearts
from P18.5, P39.5 and P90.5 were fractionated by sucrose buoyant density gradient
centrifugation. When the subcellular fractions were analyzed by Western blot, most Ncadherin and mXinβ are present in three peaks of the gradient (I, II and III) that have
different densities (representative P39.5 profile shown in Figure 4.5 A). We found an
increasing proportion of N-cadherin in the peak I as the age of the mice increases from
P18.5 to P90.5 (Figure 4.5 B), which strongly suggests that the peak I may contain the
subcellular fraction of mature ICDs. Previous studies by electron microscopy also
indicated that mature ICDs are enriched in peak I (Colaco and Evans, 1982). Importantly,
we found that a higher proportion of mXinβ than that of N-cadherin is present in the peak
I of the gradient at all three stages (Figure 4.5 B), which further supports the observed
preferential association of mXinβ with N-cadherin at the maturing ICDs of
cardiomyocytes.
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ICD defects in mXinβ-/- hearts first appear at the time when
mXinβ is normally expressed at its peak level in the wildtype hearts
Previously, we have observed that ICD components such as N-cadherin failed to
be localized to the termini of cardiomyocytes in P16.5 mXinβ-/- hearts (Wang et al.,
2010). To determine the timing of the first appearance of such defects and its relationship
to the temporal expression profile of mXinβ in wild-type hearts, we examined the
distribution pattern of N-cadherin during the maturation of ICDs (Figure 4.6). Confocal
images of frozen sections labeled for N-cadherin showed that in the wild-type hearts at
P3.5 and P7.5 (Figure 4.6 A and C), lateral puncta of N-cadherin signal are numerous and
large clusters of N-cadherin at the termini are few. Terminal localization of N-cadherin
clearly increases after P7.5 in that at P13.5, the lateral puncta become less and large
clusters that demarcated ICDs at the termini are frequent (Figure 4.6 E). After P13.5,
lateral N-cadherin puncta continue to disappear while the terminal N-cadherin signals are
accentuated (Figure 4.6 G). In the mXinβ-/- hearts, N-cadherin staining pattern is initially
indistinguishable from that of the wild-type hearts at P3.5 and P7.5 (Figure 4.6 B and D),
but in the mutant hearts at later stages, lateral puncta of N-cadherin remain numerous and
ICD-like structures are much less frequent (Figure 4.6 F, and H). The maturation process
of ICDs is diagramed in Figure 6 I, based on our previous observation of N-cadherin
distribution in embryonic hearts (Sinn et al., 2002) and the current study on the postnatal
process.
To confirm the apparent onset of the defects in the distribution N-cadherin after
P7.5, we quantified the incorporation of N-cadherin into the termini of cardiomyocytes
from P3.5 to P60.5 in the confocal images (Figure 4.6 J). The ratios of fluorescence
signals located at the termini of the cardiomyocytes versus the total signals in the entire
cardiomyocytes were calculated from confocal images of frozen sections of each
developmental stage. In the wild-type hearts, the terminal distribution of N-cadherin
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continuously increases during the developmental stages studied until P60.5 (Figure 4.6 J,
blue bars), consistent with previously reported time course of ICD maturation (Angst et
al., 1997). In contrast, the localization of N-cadherin to the termini proceeds slowly in the
mXinβ-null hearts (Figure 4.6 J, orange bars). By P13.5, statistically significant
differences of the terminal localization of N-cadherin between wild-type and mXinβ-/hearts were observed. These observations suggest the redistributions of intercellular
junctions proceed quickly between P3.5 and P13.5 and mXinβ plays indispensible roles
during this period. Without mXinβ, N-cadherin fails to be restricted to the termini of
cardiomyocytes by the time when mXinβ is expressed at its peak level in the wild-type
hearts.
Desmosomes and gap junctions also fail to be restricted to
the termini of cardiomyocytes in mXinβ-/- hearts
During the establishment and maturation of intercellular junctions in various
tissues, the adherens junctions have a leading role in determining the distribution of
desmosomes and gap junctions (Green et al., 2010; Hertig et al., 1996a; Hertig et al.,
1996b). Thus, we asked if mXinβ-/- hearts have corresponding defects in the distributions
of desmosomes and gap junctions. Quantification of the incorporation of desmoplakin (a
desmosome marker) and connexin 43 (a gap junction marker) in confocal images of
developing hearts was carried out (Figure 4.7). The terminal distribution of desmoplakin
showed a trend (Figure 4.7 A) similar to that of N-cadherin between P3.5 and P24.5 in
the wild-type hearts (Figure 4.6 J). Accordingly, a significantly lower proportion of
desmoplakin in the mXinβ-/- cardiomyocytes’ termini was observed at P13.5 (Figure 4.7
A). On the other hand, the terminal distribution of connexin 43 increases abruptly
between P13.5 and P15.5 in the wild-type hearts, consistent with the reported lag of gap
junction’s re-distribution during ICD maturation (Angst et al., 1997; Hirschy et al.,
2006), and the defect of connexin 43 localization in the mXinβ-/- hearts becomes
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significant at P15.5 (Figure 4.7 B). These data suggest that mXinβ-/- hearts have a general
defect in the developmental re-organization of all three types of intercellular junctions,
resulting in failure in the maturation of ICDs.
Intercellular junction components retain their close spatial
relationship in mXinβ-/- hearts despite being mis-localized
To determine if the failure to restrict intercellular junctions to the termini of
cardiomyocytes in the mXinβ-/- hearts could be a result of the association between
intercellular junctions normally found in the mature ICDs, we used double-label
immunofluorescence staining to examine the relationship between N-cadherin and
desmoplakin (Figure 4.8), as well as N-cadherin and connexin 43 (Figure 4.9) in the
mXinβ-/- hearts. The results showed that co-localization between N-cadherin and
desmoplakin is indistinguishable between wild-type and mXinβ-/- hearts at both P7.5 and
P24.5 (Figure 4.8), even though at P24.5, both proteins are grossly mis-localized at the
cellular level in the mXinβ-/- hearts. Interestingly, it was noted that in the mXinβ-/- hearts
at P24.5, large clusters of N-cadherin/desmoplakin signals can occasionally be found at
the termini of some cardiomyocytes, and these clusters resemble the ICDs of the wildtype hearts (Figure 4.8 L, arrows). In addition, despite the clear defects in restricting the
intercellular junctions to the termini of cardiomyocytes at P24.5, the mutant hearts at this
stage reduces the diffused N-cadherin signals on the lateral surfaces of the
cardiomyocytes and assembles discrete clusters of N-cadherin that co-localized with
desmoplakin (Figure 4.8), and many of these lateral N-cadherin/desmoplakin clusters are
elongated and adopt a perpendicular orientation relative to the longitudinal axis of the
cardiomyocytes (Figure 4.8 L, stars). This is in contrast with the N-cadherin spots in the
P7.5 hearts, which are smaller and elongate along the longitudinal axis of cardiomyocytes
on the lateral surfaces (Figure 4.8 A – F).
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Similarly, connexin 43 also shows preserved association with N-cadherin both
before and after the localization of intercellular junctions are grossly disturbed in the
mutant hearts (Figure 4.9). Close observation showed that in the mXinβ-/- hearts at P24.5,
the numerous connexin 43 spots at the lateral surface of the cardiomyocytes almost
always have at least an N-cadherin spot nearby (Figure 4.9 L, and Figure 4.10). Since Ncadherin and connexin 43 signals do not overlap and thus it is not possible to do colocalization test, I measured the distance from each connexin 43 spot to its closest Ncadherin spot (examples of center positions of the immunofluorescence signal spots
identified by ImageJ are shown in figure 4.10 A’ and B’). No statistically significant
difference was found between such distances in wild-type and mXinβ-/- hearts at both
P7.5 and P24.5 (Figure 4.10 C, Rank sum test, n >2000, p>0.05), suggesting that in the
mXinβ-/- hearts, N-cadherin and connexin 43 retained normal spatial relationship. Thus,
the mXinβ-/- hearts seem to have numerous miniature ICD-like structures containing all
three types of intercellular junction components that are ectopically formed at the lateral
surface of cardiomyocytes. The above results are consistent with our observations with
electron microscopy, which showed that the ultrastructure of ICDs seems to be largely
preserved in P15.5 mXinβ-/- hearts and the adherens junctions, gap junctions as well as
desmosomes are all found in close proximity (Wang et al., 2010).
mXinα variants are not essential for the maturation of ICDs
Because the mXinα and mXinα-a are the prevalent mXin proteins in the P0.5 and
P30.5 (Figure 4.1 and 4.2), the unique ICD maturation defect in mXinβ-/- hearts and the
absence of apparent ICD defects in the mXinα-/- hearts from young animals strongly
suggest that mXinα variants are not essential for ICD maturation. To further test this idea,
I generated mXinα-/-:mXinβ-/- double knockout (DKO) animals and studied their ICDs
by immunostaining (Figure 4.11) and Western blot (Figure 4.12). The loss of both mXinα
and mXinβ expression was confirmed by immunostaining (Figure 4.11 E) and Western
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blot (Figure 4.12) with an antibody common to both mXinα and mXinβ (U1013). I found
that loss both mXinα and mXinβ leads to ICD defects indistinguishable from the defects
caused by loss of mXinβ alone (compare Figure 4.11 to Figure 4.8 and 4.9). In the P19.5
DKO hearts, the N-cadherin, desmoplakin and connexin 43 are scattered as discrete spots
on the surface of cardiomyocytes (Figure 4.11 G – I and J – L). Similar to the mXinβ-/hearts, the ICD-like structures in the DKO hearts show apparently normal co-localization
between N-cadherin and desmoplakin, as well as normal association between N-cadherin
and connexin 43. Furthermore, Western blot analysis showed no difference in the
expression of N-cadherin, desmoplakin and connexin 43 in P13.5 wild-type, mXinα-/-,
mXinβ-/- and DKO hearts (Figure 4.12). Thus, mXinα variants are not essential for the
maturation of ICDs and further loss of mXinα variants in the mXinβ-/- hearts does not
contribute to more severe defects in the maturation of ICDs.
Interestingly, mXinβ is apparently down-regulated in the mXinα-/- heart at P13.5
(Figure 4.12 lane 2 mXinβ). We further confirmed and extended this observation by
quantitative Western blot experiments. We found that the levels of mXinβ are
significantly lower at P3.5 and P7.5 in the mXinα-/- hearts than in the wild-type hearts
(wild-type data are from figure 4.1). However, at P30.5 the level of mXinβ in mXinα-/hearts returns to normal (Figure 4.13). Thus, mXinα may play a role in maintaining
mXinβ level between P3.5 and P13.5 but not at P30.5.
Discussion
In this study, we demonstrated that mXinβ promotes ICD maturation by
promoting the restricted localization of intercellular junctions to the longitudinal termini
of cardiomyocytes during postnatal maturation of the hearts, thus allowing the
establishment of the adult arrangement of intercellular junctions between
cardiomyocytes. On the other hand, mXinβ is neither required for colocalizing of Ncadherin and desmoplakin nor targeting of connexin 43 to the vicinity of N-cadherin. We
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also showed that mXinα variants are not essential for the initial maturation of ICDs,
likely because they lack specific protein functions that are required for this process.
mXinβ plays important roles in the maturation of ICDs
We provided several lines of evidence to establish mXinβ’s critical roles in the
postnatal reorganization of intercellular junctions that leads to the restricted localization
of ICD components to the termini of cardiomyocytes.
Expression pattern of mXinβ strongly correlates with the
timing of ICD maturation
The first line of evidence is the strong correlation between the timing of ICD
maturation and mXinβ’s unique temporal expression profile. By immunofluorescence
staining for intercellular junction markers and confocal microscopy, we provided a
detailed description of the process of ICD maturation from P3.5 to P60.5 (Figure 4.6 and
4.7). Our findings are in agreement with previously reported observations that in
mammals, ICD maturation is a postnatal process, during which the three types of
intercellular junctions found in adult ICDs redistribute from the entire surface of
cardiomyocytes to the longitudinal termini of these cells (Angst et al., 1997; Hirschy et
al., 2006; Peters et al., 1994). Consistent with previous studies, we showed that
maturation of ICDs takes more than a month in rodents; as quantification showed Ncadherin localization to the cardiomyocytes’ termini continue to increase significantly
from P24.5 to P60.5 (Figure 4.6). We also confirmed that the time courses of
incorporating adherens junctions (N-cadherin as the marker) and desmosomes
(desmoplakin as the marker) to the longitudinal termini of cardiomyocytes are similar;
meanwhile, incorporation of gap junctions (connexin 43 as the marker) lags behind the
two types of adhering junctions. More importantly, we provided novel insights for the
time course of ICD maturation by providing more time points of observations between
P3.5 and P24.5 and showed that maturation of ICDs proceeds most rapidly between P3.5
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and P13.5. Indeed, the percentage of terminally localized N-cadherin increased by 3.3
fold in 10 days (from P3.5 to P13.5), whereas the next comparable degree of increase (2.3
fold) happen in 47 days (from P13.5 to P60.5). This detailed description of the time
course of ICD maturation allowed us to establish a strong correlation between the timing
of ICD maturation and the temporal expression profile of mXinβ.
The temporal expression profile of mXinβ was established by quantitative
Western blot (Figure 4.1). This technique allowed us to measure the absolute quantity of
mXinβ protein accurately, which in turn makes it possible to compare the expression of
mXinβ among different time points and to that of other proteins. We showed that the
expression of mXinβ at the protein level is dynamically and uniquely regulated. The
concentration of mXinβ increases rapidly from P0.5 to P13.5 and then falls sharply until
P20.5, which is then followed by a gradual increase that continues until at least P60.5.
Significantly, we found that the very rapid increase of mXinβ concentration between P0.5
and P13.5 correlates very well with the sharp increase of the terminal localization of Ncadherin and desmoplakin (Figure 4.6 and 4.7), supporting mXinβ may have important
roles in this initial phase of postnatal ICD maturation.
mXinβ is preferentially targeted to the maturing ICDs
The second line of evidence supporting mXinβ’s role in ICD maturation is that
during this process, mXinβ is preferentially targeted to the maturing ICDs at the termini
of cardiomyocytes where it co-localizes with N-cadherin (Figure 4.3). Such preferential
localization is unique to mXinβ because the mXinα variants were found to be colocalized with N-cadherin both at the termini and the lateral surface of the
cardiomyocytes (Figure 4.4). Although we did observe a small amount of mXinβ signals
at the lateral surface that are co-localized with bright N-cadherin puncta, majority of Ncadherin puncta at the lateral surface do not have detectable level of mXinβ. The
preferential association of mXinβ with a subpopulation of N-cadherin-containing
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complexes is further supported by subcellular fractionation experiments (Figure 4.5),
which revealed that mXinβ is preferentially associated with the mature ICD-containing
fraction during development. The preferential association between mXinβ with the
maturing ICDs suggests that mXinβ may be directly involved in the maturation of ICDs.
In the mXinβ-null hearts, ICD defects appear when mXinβ
expression reaches peak level in the wild-type hearts
The third line of evidence supporting a direct involvement of mXinβ in localizing
the intercellular junctions to the termini of cardiomyocytes is that loss of mXinβ leads to
failure of restricting all three types of intercellular junctions to the termini of
cardiomyocytes, and the onset of such defect correlates very well with the peak
expression of mXinβ in wild-type hearts (Figure 4.6 and 4.7). Previously, we have shown
that in the mXinβ-/- hearts at P16.5, the N-cadherin signal is scattered as discrete spots
across the surface of cardiomyocytes (Wang et al., 2010). However, at that time, it was
not clear when such defect occurs in the course ICD maturation. In this study, we showed
that the significantly lower percentages of the terminal localization of N-cadherin and
desmoplakin in the mXinβ-/- hearts than those in the wild-type hearts occur at P13.5. On
the other hand, a significant difference for connexin 43 localization was not observed
until P15.5, consistent with the delay in incorporating the connexin 43 to the termini of
cardiomyocytes. Thus, mXinβ is rapidly up-regulated from P0.5 and reaches its peak
expression at P13.5 while loss of mXinβ during this period leads to failure of restricting
the intercellular junction to the termini of cardiomyocytes. Taken together, the correlation
between mXinβ’s temporal expression pattern with the time course of ICD maturation,
the preferential targeting of mXinβ to the maturing ICDs, and the timing of the onset of
the defects in ICD maturation when mXinβ is lost, all point to a direct involvement of
mXinβ in this process.
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Molecular mechanisms of ICD maturation and mXinβ’s
function in this process
As diagrammed in Figure 4.6I, the components of adherens junctions seem to
undergo two phases of redistribution in order for ICDs to form. The first phase happens
during embryonic development and early postnatal stages, in which the diffusely
distributed adherens junction components aggregate to form discrete spots. The process
of forming brightly stained spots and the reduction of diffusely-stained N-cadherin on the
lateral surface of cardiomyocytes might be mediated by cadherin clustering. mXinβ-null
hearts show no apparent defects in this phase (Figure 4.6) and thus this process will not
be further discussed.
In the second phase, the clusters of adherens junction redistribute to the termini of
cardiomyocytes; specifically, the contacts at the cell termini expand to form mature ICDs
while the lateral clusters reduce both in number and in signal intensity. mXinβ-null hearts
show severe defects in this process, which results in the failure of restricting clusters of
the adherens junctions to the termini of the cells. Thus, the following discussion will
focus on potential mechanisms for the redistribution of adherens junction during ICD
maturation. Although little is known about such mechanisms in the cardiomyocytes,
principles learned from other cellular models may be valuable for our understanding of
ICD maturation because cadherin mediated cell-cell interaction is fundamental in
multicellular organisms.
The redistribution of adherens junctions might be regulated
by the stability of the junctions at different cellular sites
In epithelial cells, it has been shown that E-cadherin clusters are highly dynamic
structures. New E-cadherin molecules are continuously added to the clusters, and ATPdependent mechanisms maintains the size of clusters by actively removing E-cadherin
molecules from the cluster (Hong et al., 2010). Depletion of ATP in the cells leads to
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very large cadherin clusters, suggesting cadherin cluster size is a result of dynamic
equilibrium between addition and removal of junctional components (Hong et al., 2010).
If similar mechanisms exist for the N-cadherin clusters in the heart, they could have
important implication for the maturation of ICDs. In principle, tipping the balance
between addition and removal of N-cadherin into and from the clusters could account for
both the reduction of N-cadherin clusters at the lateral surface of cardiomyocytes and the
expansion of adherens junctions at the cell termini. Supporting this idea, cadherin
addition and removal have been shown to be critical for epithelial junction formation and
remodeling (Classen et al., 2005; Lock and Stow, 2005).
One way to control the balance between addition and removal of junctional
components is to regulate the interaction between adherens junctions and their
underlining actin cytoskeleton, which plays important roles for the stability of adherens
junctions (Green et al., 2010). Although it has been shown that a direct and stable link
between the cadherin-β-catenin complex and the actin filaments mediated solely by αcatenin is unlikely (Drees et al., 2005; Yamada et al., 2005), emerging evidence indicates
that molecules other than or in addition to α-catenin can couple the cadherin complex to
the underlining actin filaments stably, and these molecules play instrumental role for the
stability of adherens junctions. Two recent examples supporting this idea are briefly
described here: Cavey and colleagues showed that in Drosophila embryonic epithelial
cells, the spot adherens junctions (SAJs) mediated by DE-cadherin is tightly coupled to
very stable, actin depolymerizing drug-resistant actin patches. Since SAJ’s stability is αcatenin independent, Cavey et al. suggested that a factor “X” is responsible for
connecting SAJs to the actin patches (Cavey and Lecuit, 2009; Cavey et al., 2008).
Similarly, Abe and Takeichi showed that in mammalian epithelial cells, EPLIN directly
couples the E-cadherin-β-catenin complex to actin filaments in an α-catenin-dependent
fashion, and depletion of EPLIN leads to failure of forming adhesion belts (Abe and
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Takeichi, 2008). As will be discussed below, the mXin proteins might be the cardiac
counterpart of the factor X and EPLIN.
Active expansion of intercellular contacts may also play
important roles in the maturation of ICDs at the termini of
cardiomyocytes
In addition to regulating adherens junction stability at lateral surface and maturing
ICDs, the cardiomyocytes likely promote adherens junction expansion at the maturing
ICDs through active mechanisms. In the epithelial cells, the Rho family small GTPases
have been shown not only to mediate signals initiated by cadherin engagement (Perez et
al., 2008) but also to drive adherens junction initiation and expansion (Yamada and
Nelson, 2007). Rac activity promotes lamellapodia formation, which promotes contact
formation and expansion whereas RhoA also regulates contact expansion through
regulating the actomyosin contractility underlining the forming adherens junctions
(Yamada and Nelson, 2007). It has been shown in the non-muscle cells, the small
GTPase Rap1 acts upstream of both Rac and Rho to regulate adherens junctions
(Kooistra et al., 2007; Pannekoek et al., 2009). Interestingly, the Rap1 mediated signals
have been shown to enhance adherens junction formation followed by gap junction
establishment in cardiomyocytes (Somekawa et al., 2005), suggesting the roles of these
small GTPases are likely conserved in cardiomyocytes.
mXinβ may stabilize adherens junctions preferentially at
the termini of cardiomyocytes by linking the adherens
junctions to the actin cytoskeleton
We have shown that mXinα can stabilize adherens junctions through its
simultaneous and direct interacts with β-catenin and actin filaments (Choi et al., 2007)
and mXinβ likely does so by similar interactions (Grosskurth et al., 2008). Therefore, the
mXin proteins may be the cardiac counterpart of the factor X and/or EPLIN, and act as a
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stable link between the adherens junctions and actin filaments. Previously we reported
that mXinβ mRNAs are preferentially concentrated at ICDs, suggesting that newly
synthesized mXinβ can be incorporated into the termini of cardiomyocytes (GustafsonWagner et al., 2007). Through its preferential localization to the cardiomyocytes’ termini,
mXinβ could increase the stability of N-cadherin clusters locally, leading to increased
accumulation of N-cadherin at the cell termini at the expense of lateral N-cadherin
clusters, and thus could promote the maturation of ICDs. Without mXinβ, the N-cadherin
clusters at the lateral side and the termini of cardiomyocytes may be equally
stable/unstable, causing failure of accumulating N-cadherin to the termini.
mXinβ may regulate Rac1 activity locally at the maturing
ICDs for ICD expansion
Previously, we observed that in the mXinβ-null hearts, Rac1 activity is
significantly down-regulated (Wang et al., 2010). Since Rac1 plays important roles in
adherens junction formation and expansion, the reduction of Rac1 could lead to defects in
expanding the adherens junctions of ICDs at the cell termini. Consistent with this
possibility, TEM observation showed that the membranes of the maturing ICDs in the
mXinβ-null hearts are smoother than those in the wild-type hearts, suggesting a depressed
Rac1-mediated membrane raffling activity in the mutant hearts (Wang et al., 2010). Our
new observation that mXinβ is preferentially localized to the maturing ICDs further
suggests that mXinβ may function locally to promote the expansion of adherens junctions
through Rac1.
Through the above mechanisms, mXinβ may directly regulate the redistribution of
adherens junctions during ICD maturation. The redistribution of adherens junctions may
in turn facilitate the redistribution of both desmosomes and gap junctions. It has been
shown that classic cadherin mediated-adherens junctions form prior to desmosomes at
newly established intercellular contacts and adherens junctions are the prerequisites for
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the formation and correct location of the desmosomes and gap junctions (Green et al.,
2010). In the heart, induced deletion of N-cadherin in adult cardiomyocytes leads to
dissolution of the entire ICDs (Kostetskii et al., 2005), further supporting the central role
of adherens junctions in maintaining the desmosomes and gap junctions. Thus, by
regulating the localization of adherens junctions, mXinβ could control of the organization
of the desmosomes and gap junctions indirectly.
Regulation of the expression and localization of mXinβ
The expression and localization of mXinβ likely plays central roles in the
maturation of ICDs. Our previous studies showed that both the cXin and mXin are under
the control of MEF2 transcription factors (Lin et al., 2005; Wang et al., 1999). More
recent study from another group also supports this observation (Huang et al., 2006).
However, in the postnatal stage, MEF2 activity is high from birth to at least 3-week of
age in the mouse (Kolodziejczyk et al., 1999), suggesting that another mechanism also
governs the dynamic change of mXinβ expression during postnatal period.
Interestingly, we showed that mXinβ protein level is highly dependent on mXinα
during the first two postnatal weeks because loss of mXinα leads to drastic downregulation in mXinβ protein level (Figure 4.12 and 4.13). However, such dependence
seems to be less important in more mature hearts as we found that the mXinβ level
returns to normal at P30.5 in mXinα-/- hearts (Figure 4.13) and becomes significantly upregulated in adult mXinα-/- hearts (Gustafson-Wagner et al., 2007). Although the
mechanism for such intriguing relationship between the mXinα and mXinβ proteins is
unknown, it is possible that the down-regulation of mXinβ in the wild-type hearts from
P13.5 to P20.5 is dictated by the reduction in levels of mXinα variants during the same
period (Figure 4.1).
On the other hand, the preferential targeting of mXinβ but not mXinα to the
maturing ICDs at the termini of cardiomyocytes might be partly explained by the fact that
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only mXinβ mRNA is specifically localized to the ICDs (Gustafson-Wagner et al., 2007).
The mechanism underlining the localization of mXinβ mRNA remains to be determined,
but localized translation of these mRNA may be important mechanism for the preferential
targeting of mXinβ proteins to the maturing/mature ICDs.
Defects of the mXinβ-null hearts provide novel insights for
ICD formation in healthy and diseased hearts
Association between intercellular junctions is independent
from the spatial distribution of the junctions at the cellular
level
Besides providing evidence for the important roles of mXinβ in ICD maturation,
our results further imply that 1) the formation of area composita by the amalgamation of
adhering junctions and 2) the localization of gap junctions to the vicinity of adhering
junctions are independent from the overall distribution of intercellular junctions at the
cellular scale. During heart development, the amalgamation of the adhering junctions
(adherens junction and desmosomes) is a prolonged process that initiates in embryonic
stage and continuous even at 3-week of age (Borrmann et al., 2006), which overlaps with
the re-distribution of the adhering junctions at the cellular level. Now we showed that
despite the disruption of the overall distribution of intercellular junctions at the cellular
level in the mXinβ-/- hearts, the amalgamation between the adhering junctions seems to
be unaffected (Figure 4.8). Similarly, the association between the adherens junctions and
the gap junctions remains unchanged in the mXinβ-/- hearts despite the extensive mislocalization of both types of junctions (Figure 4.9 – 4.10).
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Formation of multiple intercalated discs in diseased hearts
might involve dys-regulation of mXinβ
Multiple intercalated discs are a pathological structure frequently found in the
hypertrophied canine and human myocardium (Laks et al., 1970; Maron and Ferrans,
1973). They are defined as two or more ICDs lying in tandem along the longitudinal axis
of a cardiomyocytes and are separated by less than 10 sarcomeres (Laks et al., 1970).
These abnormally arranged ICDs are formed between one cardiomyocytes and the
protruding processes from a neighboring cardiomyocyte (Maron and Ferrans, 1973).
Maron and co-workers postulated that these cellular processes are established by sidewise
addition of sarcomeres between two lateral intercellular junctions. Lateral intercellular
junctions are prominent between cardiomyocytes in embryonic and neonatal hearts, as
shown in this and previous studies on ICD development, but greatly reduce during the
maturation of the ICDs in an mXinβ-depend fashion. Thus, it is possible that dysregulation of mXinβ or its related cellular pathways may play important roles in forming
the pathological structure, multiple intercalated discs. The miniature ICD-like structures
arranged in tandem in the p24.5 mXinβ-/- hearts further supports this possibility (Figure
4.8 and 4.9).
mXinα in the maturation and maintenance of ICDs
Besides demonstrating the indispensible roles of mXinβ in postnatal maturation of
ICDs, our results also indicate that mXinα variants are not essential for the maturation of
ICDs, and this is likely due to their lack of specific protein functions for this process.
This notion is supported by the fact that although mXinα variants are more highly
expressed than mXinβ during the course of ICD development (Figure 4.1 and 4.2), loss
mXinα does not affect the maturation of ICDs (Gustafson-Wagner et al., 2007). More
importantly, in the mXinβ-/- background, additional loss of mXinα does not contribute to
more severe defects in ICD maturation (Figure 4.11). On the other hand, despite its lack
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of important roles in ICD maturation, mXinα is required for the maintenance of ICDs in
adult heart (Gustafson-Wagner et al., 2007). Interestingly, we have observed an increase
in mXinβ expression after P20.5 in the wild-type hearts; in the P60.5 hearts, mXinβ
accounts for about 1/3 of the total mXin proteins. This phenomenon indicates mXinβ
may also play important roles in the ICDs of the adult hearts.
Conclusion
In summary, we have shown that mXinβ is required for the reorganization of
intercellular junctions to establish mature ICDs in postnatal cardiomyocytes. mXinβ
likely carries out this role by nucleating the formation of mature ICDs at the termini of
cardiomyocytes and preventing the retention of intercellular contacts and formation of
ectopic ICDs at the lateral surfaces. We also showed that the overall organization of
intercellular junctions in the adult heart is neither required for the amalgamation of
adhering junctions to form the area composita nor for the association of gap junctions
and the adhering junctions. Finally, we provided evidence to show that mXinα are not
essential for the maturation of ICDs.
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Figure 4.1. Temporal expression profiles of mXinβ, mXinα variants and N-cadherin in
developing postnatal hearts established by quantitative Western blot. The amounts of the
proteins per mg of heart (A, C and D) and in the entire heart (B, D and F) are plotted
against age to establish their developmental expression profiles. mXinβ (A and B);
mXinα variants (C and D); N-cadherin (E and F). Each point represents 3 independent
heart samples. Error bars represent standard errors.
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Figure 4.2. Comparison of the expressions of mXin proteins with that of N-cadherin.
Data are compiled from same quantitative Western blots as in Figure 4.1.
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Figure 4.3. Characterization of the co-localization between mXinβ and N-cadherin during
postnatal heart development. Confocal images of double-immunofluorescence labeled
frozen sections of wild-type hearts for N-cadherin (A and D) and mXinβ (B and E). (A –
C) P7.5 and (D – F) P24.5. Merged imaged are also shown (C and F). Bar = 20 µm.
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Figure 4.4. Characterization of the co-localization between mXinα and N-cadherin during
postnatal heart development. Confocal images of double-immunofluorescence labeled
frozen sections of wild-type hearts for N-cadherin (A and D) and mXinα (B and E). (A –
C) P7.5 and (D – F) P24.5. Merged imaged are also shown (C and F). Bar = 20 µm.
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Figure 4.5. Subcellular fractionation provided evidence for the preferential association of
mXinβ with the maturing/matured ICDs. (A) Representative profiles (P39.5 hearts) of the
distributions of N-cadherin and mXinβ in the sucrose gradient. The N-cadherin is clearly
present in three distinct parts of the gradient, designated as peak I, II and III. mXinβ is
concentrated in peak I. (B) Percentage of N-cadherin and mXinβ distributed in each peak
at different developmental stages. A developmental increase of N-cadherin in the peak I
indicates that peak I contains the mature ICDs. At any given stage, larger proportion of
mXinβ than N-cadherin is associated with peak I.
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Figure 4.6. Time courses of ICD maturation in the postnatal wild-type and mXinβ-/hearts characterized by N-cadherin localization. (A – H) Confocal images of frozen
sections labeled for N-cadherin in wild-type (A, C, E, G and I) and mXinβ-/- (B, D, F, H
and J) hearts. Ages of the mice are shown at the left side of the images. Bar = 15 µm. (I)
Diagram of N-cadherin localization in the cardiomyocytes from embryonic stage to P24.5
and older. Cardiomyocytes’ surfaces drawn with grey lines and the N-cadherin
immunofluorescence signals were shown in red. (J) Quantification of the ratios of
terminally localized N-cadherin signals in wild-type and mXinβ-/- hearts.
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Figure 4.7. Characterization of the distributions of desmosome and gap junctions in the
postnatal wild-type and mXinβ-/- hearts. Quantification of the ratios of terminally
localized desmoplakin (A) and connexin 43 (B) signals in wild-type and mXinβ-/- hearts.
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Figure 4.8. Confocal images of double labeled frozen sections demonstrating the
preserved co-localization between N-cadherin and desmoplakin in the mXinβ-/- hearts.
(A, D, G and J) N-cadherin (green), (B, E, H and K) desmoplakin (magenta) and merged
images (C, F I and L). Age and genotypes are labeled at the left side of the images. Bar =
20 µm.
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Figure 4.9. Confocal images of double labeled frozen section demonstrating the
preserved association between N-cadherin and connexin 43 in the mXinβ-/- hearts. (A, D,
G and J) N-cadherin (green), (B, E, H and K) connexin 43 (magenta) and merged images
(C, F I and L). Age and genotypes are labeled at the left side of the images. Bar = 20 µm.
205
206
Figure 4.10. Quantification of the distances between connexin 43 and N-cadherin
immunofluorescence signal spots. Frozen sections from P24.5 wild-type (A) and mXinβ/- hearts (B) doubled labeled for N-cadherin (green) and connexin 43 (magenta) and the
positions of immunofluorescence spots located by the find maxima function of ImageJ
software from the confocal images (A’ and B’). Bar = 5 µm. (C) Box plots of the
distances between each connexin 43 immunofluorescence spot to its closest N-cadherin
spot in P7.5 and P24.5 wild-type and mXinβ-/- heart sections. No statistically significant
differences were found between the wild-type and mXinβ-/- hearts by Rank Sum tests.
N.S.: non-significant.
207
208
Figure 4.11. Confocal images of double labeled frozen sections from P19.5 wild-type (A
– C) and mXinα-/-:mXinβ-/- hearts (D – L). (A, D, G and J) N-cadherin; (B and E) total
mXin; (H) desmoplakin and (K) connexin 43. (C, F, I and L) are merged images from
their corresponding left panels. Bar = 20 µm.
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210
Figure 4.12. Western blot detection of representative intercellular junction proteins in
P13.5 wild-type (lane 1), mXinα-/- (lane 2), mXinβ-/- (lane 3) and mXinα-/-:mXinβ-/DKO hearts (lane 4). GAPDH was used as loading control.
211
212
Figure 4.13. Quantitative Western blot demonstrated that mXinβ is significantly down
regulated in mXinα-/- hearts at P3.5 and P7.5 but not at P30.5. Each bar represents 3 heart
samples. The wild-type data are the same as those in Figure 4.1 B. Error bars represent
standard errors.
213
214
CHAPTER V
SUMMARY AND FUTURE DIRECTION
Overall summary of thesis research
In this thesis, I described my efforts in characterizing the functions of the ICD
localized, Xin repeat-containing proteins both in vitro and in vivo. Through these efforts,
we bettered our understanding of how the adherens junctions at the ICDs are specialized
to withstand the contractile forces, and of the molecular mechanisms for the
establishment of ICDs. We also support the growing consensus that ICDs are intricate
organelles that carry out a range of functions including intercellular coupling, signaling
and ion channel surface expression.
The gene encoding Xin repeat containing proteins was first identified in our lab
by mRNA differentiation in the developing chicken hearts (Wang et al., 1996).
Experiments with chicken embryos suggested that the sole chicken Xin (cXin) plays an
important role in cardiac morphogenesis, particularly in chamber formation (Wang et al.,
1999). This function is further supported by the evolutionary history of the Xin-repeat
containing family of proteins: the emergence of these proteins coincides with the
emergence of true chambered hearts (Grosskurth et al., 2008). To further understand the
functions of the Xin repeat-containing proteins, we started the in vivo characterization of
the mammalian Xin by deleting the more rapidly evolved mXinα from the mouse genome
(Gustafson-Wagner et al., 2007). mXinα-deficient mice are viable and fertile,
nevertheless, they have late-onset cardiomyopathy with conduction defects and
particularly, their ICDs have structural defects in adulthood that implicate reduced
intercellular adhesion. An upregulation of the evolutionarily more conserved mXinβ in
the mXinα-deficient hearts suggests a compensatory role by mXinβ. The in vivo
characterization of mXinα opens many questions for us. My thesis research mainly
concerns two questions: 1) what are the molecular mechanisms of mXinα’s functions that
215
account for the observed phenotypes in the mXinα-deficient hearts; 2) what is the
function of mXinβ.
In the first chapter of this thesis, I reviewed our current knowledge of the cardiac
specific structure, ICDs, where the mXin proteins reside. The ICDs were first discovered
in 1866 but studies on these structures truly advanced since the 1990s. This chapter puts
my efforts in characterizing the mXin proteins into the perspective of understanding the
familiar yet underexplored ICDs.
In chapter II, I presented my contribution to the in vitro characterization of
mXinα. Together with postdoctoral fellow Sunju Choi and former graduate student
Elisabeth Gustafson-Wagner, we demonstrated that mXinα directly interacts with the
important adherens junction protein β-catenin and also binds and bundles actin filaments.
Importantly, we also showed that mXinα can interact with β-catenin and actin filaments
simultaneously, and that β-catenin facilitates the interaction between mXinα and the actin
filaments. This study allowed us to propose a model, in which mXinα is present in both
an open and close status. Interaction of mXinα with β-catenin promotes mXinα to adopt
the open status allowing it to interact with actin. Very likely, mXinα may act as a direct
link between the adherens junctions and the actin cytoskeleton, thus providing an
important means to strengthening the intercellular adhesion at the ICDs. Since mXinβ’s
Xin repeat region is also known to interact with actin, and mXinβ also has a highly
conserved β-catenin interaction domain, mXinβ likely shares the same function to link
adherens junction to the actin cytoskeleton. In this study, we have also revealed other
interaction partners of mXinα, such as KChIP2, p120-catenin, gelsolin, filamin,
indicating that mXinα might be able to organize a diverse range of molecules at the ICD.
In chapter III, I described my work in generation and characterization of the
mXinβ-/- mice. Through characterizing mXinβ-null mice, I demonstrated that mXinβ is
required for postnatal growth and survival of the animals. The hearts of mXinβ-/- animals
show ventricular septal defects and misaligned cardiomyocytes, which is consistent with
216
our hypothesis that the evolutionarily more conserved mXinβ plays an important role in
cardiac morphogenesis. I also showed that mXinβ-null hearts have diastolic dysfunction,
which may lead to heart failure that ultimately accounts for growth retardation and death
of the animals. This study also revealed signaling defects in the mutant hearts, which led
us to conjecture that mXinβ might be a scaffolding protein that organizes signaling
pathways at the highly specialized ICDs. I also found severe defects in the ICDs of the
P16.5 mXinβ-null mice, suggesting mXinβ may play a role in ICD formation.
In chapter IV, I further examined the roles of mXin proteins in the postnatal
formation of ICDs. I provided multiple lines of evidence to demonstrate that mXinβ but
not mXinα is required for ICD formation. In particular, loss of mXinβ leads to drastic
defects in the postnatal redistribution of the intercellular junctions during ICD formation.
In contrast, mXinα-null hearts form normal ICDs and additional loss of mXinα in the
mXinβ-/- background does not cause more severe defects in ICD formation than those
caused by loss of mXinβ alone. In addition, loss of mXinβ also leads to mis-localization
of mXinα, suggesting a functional hierarchy. On the other hand, mXinβ’s level is highly
dependent on mXinα during the first two to three weeks of postnatal life, indicating the
existence of an unknown interplay between the two members of the Xin repeatcontaining family of proteins. Through quantitative Western blot experiments, I also
revealed the quantitative relationship of the mXinβ, mXinα and N-cadherin, which
provides important information about the interaction among these proteins.
Conclusion and future direction
The Xin proteins are modular in nature, contain many copies of a 16-amino acid
“Xin” repeating unit, and locate at the ICDs. The Xin repeat defines a novel actin binding
domain. Together with the presence of other interacting domains for Mena/VASP,
filamin, gelsolin, etc., the Xin proteins are capable of regulating actin dynamics. The
highly conserved β-catenin-binding domain on the Xin proteins overlapped with the Xin
217
repeat region further suggest that the Xin proteins are involved in a novel mechanism in
linking and regulating the actin cytoskeleton to the N-cadherin-mediated adhesion in the
heart. The existence of a p120-catenin interacting domain distinct from the β-catenin
binding domain provides another regulatory role for the Xin proteins in signaling through
ICDs. Accumulated lines of evidence support the existence of a functional hierarchy
between mXinα and mXinβ. Hearts without mXinβ fail to form mature ICDs and result in
a mis-localization of mXinα. On the other hand, hearts without mXinα transiently downregulates mXinβ but forms ICDs. mXinα-null hearts develop late onset ultrastructural
ICD defects and mXinβ is up-regulated and localized normally to ICDs. The mXinα-null
cardiomyocytes have reduced transient outward potassium current density. It has become
apparent that mXinα interacts directly with Kv channel. These findings lead us to
hypothesize that the mXinβ initiates the formation of ICD, whereas the mXinα further
stabilizes the ICD. The molecular mechanisms by which Xin proteins function remain
unclear. Future detailed analysis of the binding/interacting domains on Xin proteins
should advance our understanding toward the mechanisms. For examples, in addition to
binding β-catenin, mXinα may interact with and recruit p120-catenin to facilitate ICD
maturation and the stability of the N-cadherin-based adhesion. The potential interactions
of Xin proteins with p120-catenin may modulate effectors such as Vav2 to regulate Rac1
and Rho activity. We have started to generate a cardiac-specific deletion mutant of p120catenin to test p120-catenin’s roles and its interplay with mXin in the hearts. The answers
to whether mXinα would directly interact with Cx43 and/or ZO-1 may provide accounts
for the gap junction remodeling (decrease in Cx43 amounts and altered its localization)
observed in human failing hearts and hearts from many animal models of
cardiomyopathy and arrhythmia.
Generation and characterization of inducible mXinβ knockout mice are also
underway, which will allow us to further examine the role of mXinβ in maintaining ICD
integrity in the adult heart. The information to be obtained may reveal the molecular
218
mechanisms underlying why adult ICD remodeling is always found in many cardiac
diseases, including cardiomyopathy and heart failure.
The mXinα-deficient mice exhibit cardiac phenotypes similar to that of human
dilated cardiomyopathy with conduction defect 2, whereas the mXinβ-null mice die
around weaning and exhibit congenital heart defects and severe growth retardation. The
future study of single nucleotide polymorphisms on CMYA1 and CMYA3 from human
populations with cardiomyopathy and conduction defects or with congenital heart defects
may potentially define these genes as disease-causing genes. It is therefore conceivable
that the knowledge gained from the roles of Xin proteins in cardiac development and
function will provide new insights for improved therapeutic strategies for human
cardiomyopathy, arrhythmias and heart failure.
219
APPENDIX A
RED/GREEN DOT PROCESSOR
' RED/GREEN DOT PROCESSOR (By Zachary Soch and Qinchuan Wang)
' V1.0.2
' Changes to speed up processing
' Used the proper variables in getting rows (instead of using just the X columns
' it uses both X and Y columns)
' V1.0.1
' CHANGES:
' + FIXED PERCENTAGE COMPLETE
' + IMPROVED EXCEL FREEZING PREVENTION
' Sheet2.Activate
startTime = Timer
Sheet2.Cells.Clear
Application.Calculation = xlCalculationManual
' CHANGE THIS NUMBER TO LOWER IF EXCEL FREEZES AND YOU CAN NO
LONGER SEE THE PROGRESS BAR / STATUS
' CHANGE IT IN INCREMENTS OF 250. DO NOT SET BELOW 1.
CALCULATIONS_WAIT = 6500
220
'On Error Resume Next
Dim redCoordColumn, greenCoordColumn, distanceColumn As String
Dim redXSource, redYSource, greenXSource, greenYSource As String
' Set the starting cell range (A2, B2, F2, E2, etc) as the starting point for
' each of the point sources
redXSource = "B2"
redYSource = "C2"
greenXSource = "F2"
greenYSource = "G2"
' Set the starting points for the coord column
redCoordColumn = "A2"
greenCoordColumn = "B2"
distanceColumn = "C2"
timeComplColumn = "E2"
' Create Labels in Sheet 2
Sheet2.Range(redCoordColumn).Offset(-1).Value2 = "Red Coord"
Sheet2.Range(greenCoordColumn).Offset(-1).Value2 = "Green Coord"
Sheet2.Range(distanceColumn).Offset(-1).Value2 = "Distance"
Sheet2.Range(timeComplColumn).Offset(-1).Value2 = "Completion Time"
' Loop through each cell
numRedXRows = Sheet1.Range(redXSource,
Sheet1.Range(redXSource).End(xlDown)).Rows.Count
221
numRedYRows = Sheet1.Range(redYSource,
Sheet1.Range(redYSource).End(xlDown)).Rows.Count
numGreenYRows = Sheet1.Range(greenYSource,
Sheet1.Range(greenYSource).End(xlDown)).Rows.Count
numGreenXRows = Sheet1.Range(greenXSource,
Sheet1.Range(greenXSource).End(xlDown)).Rows.Count
' Check that the number of RedXRows and RedYRows is the same
' also do the same with green dots
If numRedXRows <> numRedYRows Then
MsgBox "Please check X / Y Values for Red, there appears to be a null value in one
of the points"
Exit Sub
ElseIf numGreenYRows <> numGreenXRows Then
MsgBox "Please check X/ Y Values for Green, there appears to be a null value in
one of the points"
Exit Sub
End If
' Calculation of the percentage complete
percentCompl = 0
numCalcs = numRedXRows * numGreenXRows
numCalcsLeft = numCalcs
' Loop each red x row
For i = 0 To numRedXRows - 1
Dim ref_gxPoint, ref_gyPoint As Variant
222
' Get the rx and ry points
rxPoint = CDbl(Sheet1.Range(redXSource).Offset(i, 0).Value2)
ryPoint = CDbl(Sheet1.Range(redYSource).Offset(i, 0).Value2)
shortestDistance = ""
' Loop through the green rows for every red row
For n = 0 To numGreenXRows - 1
' set the gx,gy points
gxPoint = CDbl(Sheet1.Range(greenXSource).Offset(n, 0).Value2)
gyPoint = CDbl(Sheet1.Range(greenYSource).Offset(n, 0).Value2)
' calculate the distance (no sqrt yet!)
' this speeds the calcs up pretty quicky.
distanceX = (rxPoint - gxPoint) ^ 2
distanceY = (ryPoint - gyPoint) ^ 2
distanceC = (distanceX + distanceY)
'Sheet2.Range(distanceColumn).Offset(0, 2).Value2 = distance
' Check for shortest distance
If shortestDistance = "" Then
shortestDistance = distanceC
ref_gxPoint = gxPoint
ref_gyPoint = gyPoint
223
Else
If distanceC < shortestDistance Then
shortestDistance = distanceC
ref_gxPoint = gxPoint
ref_gyPoint = gyPoint
End If
End If
numCalcsLeft = numCalcsLeft - 1
If numCalcsLeft Mod (numCalcs * 0.01) = 0 Then
percentCompl = 100 * (numCalcs - numCalcsLeft) / numCalcs
Application.StatusBar = Round(percentCompl, 2) & "% completed. [" &
numCalcsLeft & " iterations left.]"
End If
' This prevents excel from freezing...
If CALCULATIONS_WAIT > 0 Then
If numCalcsLeft Mod CALCULATIONS_WAIT = 0 And numCalcsLeft <> 0
Then
Application.Wait (Now + TimeValue("0:00:01"))
End If
End If
Next
' Setup the new data in sheet 2
224
Sheet2.Range(redCoordColumn).Offset(i, 0).Value2 = CStr("(" & rxPoint & ", " &
ryPoint & ")")
Sheet2.Range(greenCoordColumn).Offset(i, 0).Value2 = CStr("(" & ref_gxPoint &
", " & ref_gyPoint & ")")
Sheet2.Range(distanceColumn).Offset(i, 0).Value2 = Sqr(shortestDistance)
Next
Application.Calculation = xlCalculationAutomatic
endTime = Timer
Sheet2.Range(timeComplColumn).Value2 = Format(endTime - startTime, "Fixed") &
" seconds."
225
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