* Your assessment is very important for improving the workof artificial intelligence, which forms the content of this project
Download The intercalated disc-associated Xin family of proteins in cardiac
Survey
Document related concepts
Cellular differentiation wikipedia , lookup
Magnesium transporter wikipedia , lookup
Extracellular matrix wikipedia , lookup
Cytoplasmic streaming wikipedia , lookup
Hedgehog signaling pathway wikipedia , lookup
Protein moonlighting wikipedia , lookup
Gap junction wikipedia , lookup
Cytokinesis wikipedia , lookup
Signal transduction wikipedia , lookup
List of types of proteins wikipedia , lookup
Secreted frizzled-related protein 1 wikipedia , lookup
VLDL receptor wikipedia , lookup
Transcript
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. Follow this and additional works at: http://ir.uiowa.edu/etd Part of the Biology Commons 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. 67 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. 70 71 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. 72 73 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). 74 75 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). 76 77 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. 78 79 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. 80 81 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. 82 83 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. 84 85 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. 86 87 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. 88 89 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. 90 - 91 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. 92 93 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 134 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). 94 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. 95 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 96 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). 97 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 98 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. 99 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 100 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 101 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 102 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 103 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 104 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 105 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 106 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, 10 107 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 108 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 109 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 110 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 111 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. 112 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 113 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 114 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. 115 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 116 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 117 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 118 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. 119 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). 120 121 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. 122 123 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. 124 125 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 126 127 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. 128 129 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. 130 131 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. 132 133 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 134 135 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. 136 137 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. 138 139 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). 140 141 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. 142 143 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. 144 145 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. 146 147 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 (*). 148 149 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 150 151 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. 152 153 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 182 152 153 117 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. 154 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) 155 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 1m 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) 156 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 158 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. 159 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 160 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 161 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. 162 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 163 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 37 164 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). 165 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 166 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. 167 (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). 168 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 169 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 170 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 171 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. 172 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 173 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 174 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). 175 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 176 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 177 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 P3.5 178 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 179 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. 180 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 181 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 182 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 183 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 184 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 185 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). 186 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 187 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. 188 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. 189 190 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. 191 192 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. 193 194 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. 195 196 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. 197 198 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. 199 200 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. 201 202 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. 203 204 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. 209 210 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 REFERENCES Abascal, F., Zardoya, R., and Posada, D. (2005). ProtTest: selection of best-fit models of protein evolution. Bioinformatics 21, 2104-2105. Abe, K., and Takeichi, M. (2008). EPLIN mediates linkage of the cadherin catenin complex to F-actin and stabilizes the circumferential actin belt. Proc Natl Acad Sci U S A 105, 13-19. Ai, D., Fu, X., Wang, J., Lu, M.F., Chen, L., Baldini, A., Klein, W.H., and Martin, J.F. (2007). Canonical Wnt signaling functions in second heart field to promote right ventricular growth. Proc Natl Acad Sci U S A 104, 9319-9324. Ai, Z., Fischer, A., Spray, D.C., Brown, A.M., and Fishman, G.I. (2000). Wnt-1 regulation of connexin43 in cardiac myocytes. J Clin Invest 105, 161-171. Angst, B.D., Khan, L.U., Severs, N.J., Whitely, K., Rothery, S., Thompson, R.P., Magee, A.I., and Gourdie, R.G. (1997). Dissociated spatial patterning of gap junctions and cell adhesion junctions during postnatal differentiation of ventricular myocardium. Circ Res 80, 88-94. Arulanandam, R., Vultur, A., Cao, J., Carefoot, E., Elliott, B.E., Truesdell, P.F., Larue, L., Feracci, H., and Raptis, L. (2009). Cadherin-cadherin engagement promotes cell survival via Rac1/Cdc42 and signal transducer and activator of transcription3. Mol Cancer Res 7, 1310-1327. Asimaki, A., Syrris, P., Wichter, T., Matthias, P., Saffitz, J.E., and McKenna, W.J. (2007). A novel dominant mutation in plakoglobin causes arrhythmogenic right ventricular cardiomyopathy. Am J Hum Genet 81, 964-973. Barbato, J.C., Huang, Q.Q., Hossain, M.M., Bond, M., and Jin, J.P. (2005). Proteolytic N-terminal truncation of cardiac troponin I enhances ventricular diastolic function. J Biol Chem 280, 6602-6609. Barker, R.J., Price, R.L., and Gourdie, R.G. (2002). Increased association of ZO-1 with connexin43 during remodeling of cardiac gap junctions. Circ Res 90, 317-324. Basso, C., Czarnowska, E., Della Barbera, M., Bauce, B., Beffagna, G., Wlodarska, E.K., Pilichou, K., Ramondo, A., Lorenzon, A., Wozniek, O., et al. (2006). Ultrastructural evidence of intercalated disc remodelling in arrhythmogenic right ventricular cardiomyopathy: an electron microscopy investigation on endomyocardial biopsies. Eur Heart J 27, 1847-1854. Bass-Zubek, A.E., Godsel, L.M., Delmar, M., and Green, K.J. (2009). Plakophilins: multifunctional scaffolds for adhesion and signaling. Curr Opin Cell Biol 21, 708716. Baurand, A., Zelarayan, L., Betney, R., Gehrke, C., Dunger, S., Noack, C., Busjahn, A., Huelsken, J., Taketo, M.M., Birchmeier, W., et al. (2007). Beta-catenin downregulation is required for adaptive cardiac remodeling. Circ Res 100, 13531362. 226 Behrens, J., Jerchow, B.-A., Wurtele, M., Grimm, J., Asbrand, C., Wirtz, R., Kuhl, M., Wedlich, D., and Birchmeier, W. (1998). Functional interaction of an axin homolog, conductin, with b-catenin, APC, and GSK3b. Science 280, 586-599. Behrens, J., von Kries, J.P., Kuhl, M., Bruhn, L., Wedlich, D., Grosschedl, R., and Birchmeier, W. (1996). Functional interaction of β-catenin with the transcription factor LEF-1. Nature 382, 638-642. Bennett, P.M., Maggs, A.M., Baines, A.J., and Pinder, J.C. (2006). The transitional junction: a new functional subcellular domain at the intercalated disc. Mol Biol Cell 17, 2091-2100. Bierkamp, C., McLaughlin, K.J., Schwarz, H., Huber, O., and Kemler, R. (1996). Embryonic heart and skin defects in mice lacking plakoglobin. Dev Biol 180, 780-785. Boengler, K., Hilfiker-Kleiner, D., Drexler, H., Heusch, G., and Schulz, R. (2008). The myocardial JAK/STAT pathway: from protection to failure. Pharmacol Ther 120, 172-185. Bonne, S., Gilbert, B., Hatzfeld, M., Chen, X., Green, K.J., and van Roy, F. (2003). Defining desmosomal plakophilin-3 interactions. J Cell Biol 161, 403-416. Borrmann, C.M., Grund, C., Kuhn, C., Hofmann, I., Pieperhoff, S., and Franke, W.W. (2006). The area composita of adhering junctions connecting heart muscle cells of vertebrates. II. Colocalizations of desmosomal and fascia adhaerens molecules in the intercalated disk. Eur J Cell Biol 85, 469-485. Braga, V.M., and Yap, A.S. (2005). The challenges of abundance: epithelial junctions and small GTPase signalling. Curr Opin Cell Biol 17, 466-474. Bruce, A.F., Rothery, S., Dupont, E., and Severs, N.J. (2008). Gap junction remodelling in human heart failure is associated with increased interaction of connexin43 with ZO-1. Cardiovasc Res 77, 757-765. Buckingham, M., Meilhac, S., and Zaffran, S. (2005). Building the mammalian heart from two sources of myocardial cells. Nat Rev Genet 6, 826-835. Butcher, J.T., and Markwald, R.R. (2007). Valvulogenesis: the moving target. Philos Trans R Soc Lond B Biol Sci 362, 1489-1503. Calkins, C.C., Hoepner, B.L., Law, C.M., Novak, M.R., Setzer, S.V., Hatzfeld, M., and Kowalczyk, A.P. (2003). The Armadillo family protein p0071 is a VE-cadherinand desmoplakin-binding protein. J Biol Chem 278, 1774-1783. Cavey, M., and Lecuit, T. (2009). Molecular bases of cell-cell junctions stability and dynamics. Cold Spring Harb Perspect Biol 1, a002998. Cavey, M., Rauzi, M., Lenne, P.F., and Lecuit, T. (2008). A two-tiered mechanism for stabilization and immobilization of E-cadherin. Nature 453, 751-756. 227 Chan, F.C., Cheng, C.P., Wu, K.H., Chen, Y.C., Hsu, C.H., Gustafson-Wagner, E.A., Lin, J.L., Wang, Q., Lin, J.J., and Lin, C.I. (2011). Intercalated disc-associated protein, mXin-alpha, influences surface expression of ITO currents in ventricular myocytes. Front Biosci (Elite Ed) 3, 1425-1442. Chen, S., Guttridge, D.C., You, Z., Zhang, Z., Fribley, A., Mayo, M.W., Kitajewski, J., and Wang, C.-Y. (2001). Wnt-1 signaling inhibits apoptosis by activating βcatenin/T cell factor-mediated transcription. J Cell Biol 152, 87-96. Chen, X., Bonne, S., Hatzfeld, M., van Roy, F., and Green, K.J. (2002). Protein binding and functional characterization of plakophilin 2. Evidence for its diverse roles in desmosomes and beta -catenin signaling. J Biol Chem 277, 10512-10522. Chen, X., Shevtsov, S.P., Hsich, E., Cui, L., Haq, S., Aronovitz, M., Kerkela, R., Molkentin, J.D., Liao, R., Salomon, R.N., et al. (2006). The beta-catenin/T-cell factor/lymphocyte enhancer factor signaling pathway is required for normal and stress-induced cardiac hypertrophy. Mol Cell Biol 26, 4462-4473. Cheng, C.P., Loh, Y.X., Lin, C.I., Lai, Y.J., Chen, Y.C., Sytwn, H.K., GustafsonWagner, E.A., and Lin, J.J.-C. (2005). Electrophysiological characteristics of ventricular myocytes of Xinα-deficient mice. In International Proceedings: Advances in heart disease, A. Kimchi, ed. (Bologna, Italy, MEDIMOND), pp. 2529. Cherepanova, O., Orlova, A., Galkin, V.E., van der Ven, P.F., Furst, D.O., Jin, J.P., and Egelman, E.H. (2006). Xin-repeats and nebulin-like repeats bind to F-actin in a similar manner. J Mol Biol 356, 714-723. Choi, S., Gustafson-Wagner, E.A., Wang, Q., Harlan, S.M., Sinn, H.W., Lin, J.L., and Lin, J.J. (2007). The intercalated disc protein, mXinα, is capable of interacting with β-catenin and bundling actin filaments. J Biol Chem 282, 36024-36036. Chou, P.Y. (1990). Prediction of protein structural classes from amino acid composition (New York, NY, Plenum Press). Chou, P.Y., and Fasman, G.D. (1978). Prediction of the secondary structure of proteins fromtheir amino acid sequence. Adv Enzymol 47, 45-148. Clark, K.L., Yutzey, K.E., and Benson, D.W. (2006). Transcription factors and congenital heart defects. Annu Rev Physiol 68, 97-121. Classen, A.K., Anderson, K.I., Marois, E., and Eaton, S. (2005). Hexagonal packing of Drosophila wing epithelial cells by the planar cell polarity pathway. Dev Cell 9, 805-817. Clerk, A., Pham, F.H., Fuller, S.J., Sahai, E., Aktories, K., Marais, R., Marshall, C., and Sugden, P.H. (2001). Regulation of mitogen-activated protein kinases in cardiac myocytes through the small G protein Rac1. Mol Cell Biol 21, 1173-1184. Cohen, E.D., Wang, Z., Lepore, J.J., Lu, M.M., Taketo, M.M., Epstein, D.J., and Morrisey, E.E. (2007). Wnt/beta-catenin signaling promotes expansion of Isl-1positive cardiac progenitor cells through regulation of FGF signaling. J Clin Invest 117, 1794-1804. 228 Cohen, S.A. (1996). Immunocytochemical localization of rH1 sodium channel in adult rat heart atria and ventricle. Presence in terminal intercalated disks. Circulation 94, 3083-3086. Colaco, C.A., and Evans, W.H. (1981). A biochemical dissection of the cardiac intercalated disk: isolation of subcellular fractions containing fascia adherentes and gap junctions. Journal of Cell Science 52, 313-325. Colaco, C.A., and Evans, W.H. (1982). Partial purification of an intercalated disccontaining cardiac plasma membrane fraction. Biochim Biophys Acta 684, 40-46. Coppen, S.R., Kaba, R.A., Halliday, D., Dupont, E., Skepper, J.N., Elneil, S., and Severs, N.J. (2003). Comparison of connexin expression patterns in the developing mouse heart and human foetal heart. Molecular and cellular biochemistry 242, 121-127. Cowin, P., Kapprell, H.P., Franke, W.W., Tamkun, J., and Hynes, R.O. (1986). Plakoglobin: a protein common to different kinds of intercellular adhering junctions. Cell 46, 1063-1073. Delmar, M., and McKenna, W.J. (2010). The cardiac desmosome and arrhythmogenic cardiomyopathies: from gene to disease. Circ Res 107, 700-714. Drees, F., Pokutta, S., Yamada, S., Nelson, W.J., and Weis, W.I. (2005). α-catenin is a molecular switch that binds E-cadherin-β-catenin and regulates actin-filament assembly. Cell 123, 903-915. Duka, A., Schwartz, F., Duka, I., Johns, C., Melista, E., Gavras, I., and Gavras, H. (2006). A novel gene (Cmya3) induced in the heart by angiotensin II-dependent but not salt-dependent hypertension in mice. Am J Hypertens 19, 275-281. Dupont, E., Matsushita, T., Kaba, R., Vozzi, C., Coppen, S.R., Khan, N., Kaprielian, R., Yacoub, M.H., and Severs, N.J. (2001). Altered connexin expression in human congestive heart failure. J Mol Cell Cardiol 33, 359-371. Edmondson, D.G., Lyons, G.E., Martin, J.F., and Olson, E.N. (1994). Mef2 gene expression marks the cardiac and skeletal muscle lineages during mouse embryogenesis. Development 120, 1251-1263. Eigenthaler, M., Engelhardt, S., Schinke, B., Kobsar, A., Schmitteckert, E., Gambaryan, S., Engelhardt, C.M., Krenn, V., Eliava, M., Jarchau, T., et al. (2003). Disruption of cardiac Ena-VASP protein localization in intercalated disks causes dilated cardiomyopathy. Am J Physiol Heart Circ Physiol 285, H2471-2481. Eijsbouts, S.C., Houben, R.P., Blaauw, Y., Schotten, U., and Allessie, M.A. (2004). Synergistic action of atrial dilation and sodium channel blockade on conduction in rabbit atria. J Cardiovasc Electrophysiol 15, 1453-1461. Eisenberg, L.M., and Markwald, R.R. (1995). Molecular regulation of atrioventricular valvuloseptal morphogenesis. Circ Res 77, 1-6. Eppenberger, H.M., and Zuppinger, C. (1999). In vitro reestablishment of cell-cell contacts in adult rat cardiomyocytes. Functional role of transmembrane components in the formation of new intercalated disk-like cell contacts. Faseb J 13 Suppl, S83-89. 229 Estigoy, C.B., Ponten, F., Odeberg, J., Herbert, B., Guilhuas, M., Charleston, M., Ho, J.W.K., Cameron, D., and dos Remedies, C.G. (2009). Intercalated discs:multiple proteins perform multiple functions in non-failing and failing human hearts. Biophys Rev 1, 43-49. Fentzke, R.C., Buck, S.H., Patel, J.R., Lin, H., Wolska, B.M., Stojanovic, M.O., Martin, A.F., Solaro, R.J., Moss, R.L., and Leiden, J.M. (1999). Impaired cardiomyocyte relaxation and diastolic function in transgenic mice expressing slow skeletal troponin I in the heart. J Physiol 517 ( Pt 1), 143-157. Fernandez, E., Siddiquee, Z., and Shohet, R.V. (2001). Apoptosis and proliferation in the neonatal murine heart. Dev Dyn 221, 302-310. Ferreira-Cornwell, M.C., Luo, Y., Narula, N., Lenox, J.M., Lieberman, M., and Radice, G.L. (2002). Remodeling the intercalated disc leads to cardiomyopathy in mice misexpressing cadherins in the heart. Journal of Cell Science 115, 1623-1634. Fidler, L.M., Wilson, G.J., Liu, F., Cui, X., Scherer, S.W., Taylor, G.P., and Hamilton, R.M. (2009). Abnormal connexin43 in arrhythmogenic right ventricular cardiomyopathy caused by plakophilin-2 mutations. J Cell Mol Med 13, 42194228. Forbes, M.S., and Sperelakis, N. (1985). Intercalated disc of mammalian heart: a review of structure and function. Tissue Cell 17, 605-648. Forbes, M.S., and Sperelakis, N. (1985). Intercalated discs of mammalian heart: a review of structure and function. Tissue Cell 17, 605-648. Force, T., Woulfe, K., Koch, W.J., and Kerkela, R. (2007). Molecular scaffolds regulate bidirectional crosstalk between Wnt and classical seven-transmembrane-domain receptor signaling pathways. Sci STKE 2007, pe41. Franke, W.W., Borrmann, C.M., Grund, C., and Pieperhoff, S. (2006). The area composita of adhering junctions connecting heart muscle cells of vertebrates. I. Molecular definition in intercalated disks of cardiomyocytes by immunoelectron microscopy of desmosomal proteins. Eur J Cell Biol 85, 69-82. Franke, W.W., Rickelt, S., Barth, M., and Pieperhoff, S. (2009). The junctions that don't fit the scheme: special symmetrical cell-cell junctions of their own kind. Cell Tissue Res 338, 1-17. Fukuyama, T., Ogita, H., Kawakatsu, T., Inagaki, M., and Takai, Y. (2006). Activation of Rac by cadherin through the c-Src-Rap1-phosphatidylinositol 3-kinase-Vav2 pathway. Oncogene 25, 8-19. Gallicano, G.I., Kouklis, P., Bauer, C., Yin, M., Vasioukhin, V., Degenstein, L., and Fuchs, E. (1998). Desmoplakin is required early in development for assembly of desmosomes and cytoskeletal linkage. J Cell Biol 143, 2009-2022. Garcia-Gras, E., Lombardi, R., Giocondo, M.J., Willerson, J.T., Schneider, M.D., Khoury, D.S., and Marian, A.J. (2006). Suppression of canonical Wnt/betacatenin signaling by nuclear plakoglobin recapitulates phenotype of arrhythmogenic right ventricular cardiomyopathy. J Clin Invest 116, 2012-2021. 230 Gates, J., and Peifer, M. (2005). Can 1000 reviews be wrong? actin, α-catenin, and adherens junctions. Cell 123, 769-772. Gerull, B., Heuser, A., Wichter, T., Paul, M., Basson, C.T., McDermott, D.A., Lerman, B.B., Markowitz, S.M., Ellinor, P.T., MacRae, C.A., et al. (2004). Mutations in the desmosomal protein plakophilin-2 are common in arrhythmogenic right ventricular cardiomyopathy. Nat Genet 36, 1162-1164. Giepmans, B.N. (2004). Gap junctions and connexin-interacting proteins. Cardiovasc Res 62, 233-245. Giepmans, B.N., and Moolenaar, W.H. (1998). The gap junction protein connexin43 interacts with the second PDZ domain of the zona occludens-1 protein. Curr Biol 8, 931-934. Gonsior, S.M., Guautel, M., and Hinssen, H. (1998). A six-module human nebulin fragment bundles actin filaments and induces actin polymerization. J Muscle Res Cell Motil 19, 225-235. Goossens, S., Janssens, B., Bonne, S., De Rycke, R., Braet, F., van Hengel, J., and van Roy, F. (2007). A unique and specific interaction between alphaT-catenin and plakophilin-2 in the area composita, the mixed-type junctional structure of cardiac intercalated discs. J Cell Sci 120, 2126-2136. Green, K.J., Getsios, S., Troyanovsky, S., and Godsel, L.M. (2010). Intercellular junction assembly, dynamics, and homeostasis. Cold Spring Harb Perspect Biol 2, a000125. Grosskurth, S.E., Bhattacharya, D., Wang, Q., and Lin, J.J. (2008). Emergence of Xin demarcates a key innovation in heart evolution. PLoS ONE 3, e2857. Grossmann, K.S., Grund, C., Huelsken, J., Behrend, M., Erdmann, B., Franke, W.W., and Birchmeier, W. (2004). Requirement of plakophilin 2 for heart morphogenesis and cardiac junction formation. J Cell Biol 167, 149-160. Guindon, S., and Gascuel, O. (2003). A simple, fast, and accurate algorithm to estimate large phylogenies by maximum likelihood. Syst Biol 52, 696-704. Guo, W., Li, H., Aimond, F., Johns, D.C., Rhodes, K.J., Trimmer, J.S., and Nerbonne, J.M. (2002). Role of heteromultimers in the generation of myocardial transient outward K+ currents. Circ Res 90, 586-593. Gustafson-Wagner, E.A., Sinn, H.W., Chen, Y.L., Wang, D.Z., Reiter, R.S., Lin, J.L., Yang, B., Williamson, R.A., Chen, J., Lin, C.I., et al. (2007). Loss of mXinα, an intercalated disk protein, results in cardiac hypertrophy and cardiomyopathy with conduction defects. Am J Physiol Heart Circ Physiol 293, H2680-2692. Haq, S., Michael, A., Andreucci, M., Bhattacharya, K., Dotto, P., Walters, B., Woodgett, J., Kilter, H., and Force, T. (2003). Stabilization of beta-catenin by a Wntindependent mechanism regulates cardiomyocyte growth. Proc Natl Acad Sci U S A 100, 4610-4615. Hatzfeld, M. (2005). The p120 family of cell adhesion molecules. Eur J Cell Biol 84, 205-214. 231 Hatzfeld, M., Green, K.J., and Sauter, H. (2003). Targeting of p0071 to desmosomes and adherens junctions is mediated by different protein domains. J Cell Sci 116, 12191233. Hatzfeld, M., and Nachtsheim, C. (1996). Cloning and characterization of a new armadillo family member, p0071, associated with the junctional plaque: evidence for a subfamily of closely related proteins. J Cell Sci 109 ( Pt 11), 2767-2778. Hertig, C.M., Butz, S., Koch, S., Eppenberger-Eberhardt, M., Kemler, R., and Eppenberger, H.M. (1996). N-cadherin in adult rat cardiomyocytes in culture. II. Spatio-temporal appearance of proteins involved in cell-cell contact and communication. Formation of two distinct N-cadherin/catenin complexes. Journal of Cell Science 109, 11-20. Hertig, C.M., Eppenberger-Eberhardt, M., Koch, S., and Eppenberger, H.M. (1996). Ncadherin in adult rat cardiomyocytes in culture. I. Functional role of N-cadherin and impairment of cell-cell contact by a truncated N-cadherin mutant. J Cell Sci 109 ( Pt 1), 1-10. Heuberger, J., and Birchmeier, W. (2010). Interplay of cadherin-mediated cell adhesion and canonical Wnt signaling. Cold Spring Harb Perspect Biol 2, a002915. Hill, J.A., Karimi, M., Kutschke, W., Davisson, R.L., Zimmerman, K., Wang, Z., Kerber, R.E., and Weiss, R.M. (2000). Cardiac hypertrophy is not a required compensatory response to short-term pressure overload. Circulation 101, 28632869. Hinton, R.B., Jr., Alfieri, C.M., Witt, S.A., Glascock, B.J., Khoury, P.R., Benson, D.W., and Yutzey, K.E. (2008). Mouse heart valve structure and function: echocardiographic and morphometric analyses from the fetus through the aged adult. Am J Physiol Heart Circ Physiol 294, H2480-2488. Hirakow, R., and Gotoh, T. (1980). Ontogenetic implication of the myocardial ultrastructure in the development of mammalian heart. In Etiology and morphogenesis of congenital heart disease, R. van Praagh, and A. Takao, eds. (Mount Kisco, NY, Futura Publishing Co.), pp. 99-108. Hirschy, A., Croquelois, A., Perriard, E., Schoenauer, R., Agarkova, I., Hoerstrup, S.P., Taketo, M.M., Pedrazzini, T., Perriard, J.C., and Ehler, E. (2010). Stabilised betacatenin in postnatal ventricular myocardium leads to dilated cardiomyopathy and premature death. Basic Res Cardiol. Hirschy, A., Schatzmann, F., Ehler, E., and Perriard, J.C. (2006). Establishment of cardiac cytoarchitecture in the developing mouse heart. Dev Biol 289, 430-441. Hong, S., Troyanovsky, R.B., and Troyanovsky, S.M. (2010). Spontaneous assembly and active disassembly balance adherens junction homeostasis. Proc Natl Acad Sci U S A 107, 3528-3533. Honsho, S., Nishikawa, S., Amano, K., Zen, K., Adachi, Y., Kishita, E., Matsui, A., Katsume, A., Yamaguchi, S., Nishikawa, K., et al. (2009). Pressure-mediated hypertrophy and mechanical stretch induces IL-1 release and subsequent IGF-1 generation to maintain compensative hypertrophy by affecting Akt and JNK pathways. Circ Res 105, 1149-1158. 232 Huang, H.T., Brand, O.M., Mathew, M., Ignatiou, C., Ewen, E.P., McCalmon, S.A., and Naya, F.J. (2006). Myomaxin is a novel transcriptional target of MEF2A that encodes a Xin-related alpha-actinin-interacting protein. J Biol Chem 281, 3937039379. Hunter, A.W., Barker, R.J., Zhu, C., and Gourdie, R.G. (2005). Zonula occludens-1 alters connexin43 gap junction size and organization by influencing channel accretion. Mol Biol Cell 16, 5686-5698. Ikeda, S., Kishida, S., Yamamoto, H., Murai, H., Koyama, S., and Kikuchi, A. (1998). Axin, a negative regulator of the Wnt signaling pathway, forms a complex with GSK-3β and β-catenin and promotes GSK-3β-dependent phosphorylation of βcatenin. EMBO J 17, 1371-1384. Ishiwata, T., Nakazawa, M., Pu, W.T., Tevosian, S.G., and Izumo, S. (2003). Developmental changes in ventricular diastolic function correlate with changes in ventricular myoarchitecture in normal mouse embryos. Circ Res 93, 857-865. Itoh, M., Nagafuchi, A., Moroi, S., and Tsukita, S. (1997). Involvement of ZO-1 in cadherin-based cell adhesion through its direct binding to alpha catenin and actin filaments. J Cell Biol 138, 181-192. Ji, X., Zhang, P., Armstrong, R.N., and Gilliland, G.L. (1992). The three-dimensional structure of a glutathione S-transferase from the mu gene class. Structural analysis of the binary complex of isoenzyme 3-3 and glutathione at 2.2 A resolution. Biochemistry 31, 10169-10184. Kalmyrzaev, B., Aldashev, A., Khalmatov, M., Polupanov, A., Jumagulova, A., Mamanova, L., Wilkins, M.R., and Town, M. (2006). Genome-wide scan for premature hypertension supports linkage to chromosome 2 in a large Kyrgyz family. Hypertension 48, 908-913. Kanno, M., Aoyama, Y., Isa, Y., Yamamoto, Y., and Kitajima, Y. (2008). P120 catenin is associated with desmogleins when desmosomes are assembled in high-Ca2+ medium but not when disassembled in low-Ca2+ medium in DJM-1 cells. J Dermatol 35, 317-324. Kanno, M., Isa, Y., Aoyama, Y., Yamamoto, Y., Nagai, M., Ozawa, M., and Kitajima, Y. (2008). P120-catenin is a novel desmoglein 3 interacting partner: identification of the p120-catenin association site of desmoglein 3. Exp Cell Res 314, 1683-1692. Kaplan, S.R., Gard, J.J., Carvajal-Huerta, L., Ruiz-Cabezas, J.C., Thiene, G., and Saffitz, J.E. (2004). Structural and molecular pathology of the heart in Carvajal syndrome. Cardiovasc Pathol 13, 26-32. Kaplan, S.R., Gard, J.J., Protonotarios, N., Tsatsopoulou, A., Spiliopoulou, C., Anastasakis, A., Squarcioni, C.P., McKenna, W.J., Thiene, G., Basso, C., et al. (2004). Remodeling of myocyte gap junctions in arrhythmogenic right ventricular cardiomyopathy due to a deletion in plakoglobin (Naxos disease). Heart Rhythm 1, 3-11. 233 Kaprielian, R.R., Gunning, M., Dupont, E., Sheppard, M.N., Rothery, S.M., Underwood, R., Pennell, D.J., Fox, K., Pepper, J., Poole-Wilson, P.A., et al. (1998). Downregulation of immunodetectable connexin43 and decreased gap junction size in the pathogenesis of chronic hibernation in the human left ventricle. Circulation 97, 651-660. Keil, R., Wolf, A., Huttelmaier, S., and Hatzfeld, M. (2007). Beyond regulation of cell adhesion: local control of RhoA at the cleavage furrow by the p0071 catenin. Cell Cycle 6, 122-127. Kitamura, H., Ohnishi, Y., Yoshida, A., Okajima, K., Azumi, H., Ishida, A., Galeano, E.J., Kubo, S., Hayashi, Y., Itoh, H., et al. (2002). Heterogeneous loss of connexin43 protein in nonischemic dilated cardiomyopathy with ventricular tachycardia. J Cardiovasc Electrophysiol 13, 865-870. Klaus, A., Saga, Y., Taketo, M.M., Tzahor, E., and Birchmeier, W. (2007). Distinct roles of Wnt/beta-catenin and Bmp signaling during early cardiogenesis. Proc Natl Acad Sci U S A 104, 18531-18536. Klymkowsky, M.W., Williams, B.O., Barish, G.D., Varmus, H.E., and Vourgourakis, Y.E. (1999). Membrane-anchored plakoglobins have multiple mechanisms of action in Wnt signaling. Mol Biol Cell 10, 3151-3169. Kolodziejczyk, S.M., Wang, L., Balazsi, K., DeRepentigny, Y., Kothary, R., and Megeney, L.A. (1999). MEF2 is upregulated during cardiac hypertrophy and is required for normal post-natal growth of the myocardium. Current biology : CB 9, 1203-1206. Kooistra, M.R., Dube, N., and Bos, J.L. (2007). Rap1: a key regulator in cell-cell junction formation. J Cell Sci 120, 17-22. Kostetskii, I., Li, J., Xiong, Y., Zhou, R., Ferrari, V.A., Patel, V.V., Molkentin, J.D., and Radice, G.L. (2005). Induced deletion of the N-cadherin gene in the heart leads to dissolution of the intercalated disc structure. Circ Res 96, 346-354. Kostin, S., Dammer, S., Hein, S., Klovekorn, W.P., Bauer, E.P., and Schaper, J. (2004). Connexin 43 expression and distribution in compensated and decompensated cardiac hypertrophy in patients with aortic stenosis. Cardiovasc Res 62, 426-436. Kostin, S., Rieger, M., Dammer, S., Hein, S., Richter, M., Klovekorn, W.P., Bauer, E.P., and Schaper, J. (2003). Gap junction remodeling and altered connexin43 expression in the failing human heart. Mol Cell Biochem 242, 135-144. Kucera, J.P., Rohr, S., and Rudy, Y. (2002). Localization of sodium channels in intercalated disks modulates cardiac conduction. Circ Res 91, 1176-1182. Kurdi, M., and Booz, G.W. (2007). Can the protective actions of JAK-STAT in the heart be exploited therapeutically? Parsing the regulation of interleukin-6-type cytokine signaling. J Cardiovasc Pharmacol 50, 126-141. Kwiatkowski, A.V., Weis, W.I., and Nelson, W.J. (2007). Catenins: playing both sides of the synapse. Curr Opin Cell Biol 19, 551-556. 234 Lai, Y.J., Chen, Y.Y., Cheng, C.P., Lin, J.J., Chudorodova, S.L., Roshchevskaya, I.M., Roshchevsky, M.P., Chen, Y.C., and Lin, C.I. (2007). Changes in ionic currents and reduced conduction velocity in hypertrophied ventricular myocardium of Xinα-deficient mice. Anatol J Cardiol 7 Suppl 1, 90-92. Lai, Y.J., Huang, E.Y., Yeh, H.I., Chen, Y.L., Lin, J.J., and Lin, C.I. (2008). On the mechanisms of arrhythmias in the myocardium of mXinalpha-deficient murine left atrial-pulmonary veins. Life Sci 83, 272-283. Laks, M.M., Morady, F., Adomian, G.E., and Swan, H.J. (1970). Presence of widened and multiple intercalated discs in the hypertrophied canine heart. Circ Res 27, 391-402. Lang, R.M., Bierig, M., Devereux, R.B., Flachskampf, F.A., Foster, E., Pellikka, P.A., Picard, M.H., Roman, M.J., Seward, J., Shanewise, J., et al. (2006). Recommendations for chamber quantification. Eur J Echocardiogr 7, 79-108. le Duc, Q., Shi, Q., Blonk, I., Sonnenberg, A., Wang, N., Leckband, D., and de Rooij, J. (2010). Vinculin potentiates E-cadherin mechanosensing and is recruited to actinanchored sites within adherens junctions in a myosin II-dependent manner. J Cell Biol 189, 1107-1115. Legato, M.J. (1979). Cellular mechanisms of normal growth in the mammalian heart. I. Qualitative and quantitative features of ventricular architecture in the dog from birth to five months of age. Circ Res 44, 250-262. Levy, D.E., and Darnell, J.E., Jr. (2002). Stats: transcriptional control and biological impact. Nat Rev Mol Cell Biol 3, 651-662. Li, J., Levin, M.D., Xiong, Y., Petrenko, N., Patel, V.V., and Radice, G.L. (2008). Ncadherin haploinsufficiency affects cardiac gap junctions and arrhythmic susceptibility. J Mol Cell Cardiol 44, 597-606. Li, J., Patel, V.V., Kostetskii, I., Xiong, Y., Chu, A.F., Jacobson, J.T., Yu, C., Morley, G.E., Molkentin, J.D., and Radice, G.L. (2005). Cardiac-specific loss of Ncadherin leads to alteration in connexins with conduction slowing and arrhythmogenesis. Circ Res 97, 474-481. Li, J., Patel, V.V., and Radice, G.L. (2006). Dysregulation of cell adhesion proteins and cardiac arrhythmogenesis. Clin Med Res 4, 42-52. Li, J., and Radice, G.L. (2010). A new perspective on intercalated disc organization: implications for heart disease. Dermatol Res Pract 2010, 207835. Li, J., Swope, D., Raess, N., Cheng, L., Muller, E.J., and Radice, G.L. (2011). Cardiacrestricted deletion of plakoglobin results in progressive cardiomyopathy and activation of {beta}-catenin signaling. Mol Cell Biol. Li, Y., Lin, J.J.-C., Reiter, R.S., Daniels, K., Soll, D.R., and Lin, J.J.-C. (2004). Caldesmon mutant defective in Ca2+-calmodulin binding interferes with assembly of stress fibers and affects cell morphology, growth and motility. J Cell Sci 117, 3593-3604. 235 Lin, J.J.-C., Chou, C.-S., and Lin, J.L.-C. (1985). Monoclonal antibodies against chicken tropomyosin isoforms: Production, characterization and application. Hybridoma 4, 223-242. Lin, J.J.-C., Gustafson-Wagner, E.A., Sinn, H.W., Choi, S., Jaacks, S.M., Wang, D.Z., Evans, S., and Lin, J.L.-C. (2005). Structure, expression, and function of a novel intercalated disc protein, Xin. J Med Sci 25, 215-222. Lin, J.J.-C., Wang, D.-Z., Reiter, R.S., Wang, Q., Lin, J.L.-C., and Williams, H.S. (2001). Differentially expressed genes and cardiac morphogenesis. In Formation of the Heart and Its Regulation, R.J. Tomanek, and R. Runyan, eds. (Boston, MA, Birkhauser), pp. 75-96. Lin, Q., Schwarz, J., Bucana, C., and Olson, E.N. (1997). Control of mouse cardiac morphogenesis and myogenesis by transcription factor MEF2C. Science 276, 1404-1407. Lints, T.J., Parsons, L.M., Hartley, L., Lyons, I., and Harvey, R.P. (1993). Nkx-2.5: a novel murine homeobox gene expressed in early heart progenitor cells and their myogenic descendants. Development 119, 969. Lock, J.G., and Stow, J.L. (2005). Rab11 in recycling endosomes regulates the sorting and basolateral transport of E-cadherin. Mol Biol Cell 16, 1744-1755. Lombardi, R., Dong, J., Rodriguez, G., Bell, A., Leung, T.K., Schwartz, R.J., Willerson, J.T., Brugada, R., and Marian, A.J. (2009). Genetic fate mapping identifies second heart field progenitor cells as a source of adipocytes in arrhythmogenic right ventricular cardiomyopathy. Circ Res 104, 1076-1084. Longo, K.A., Kennell, J.A., Ochocinska, M.J., Ross, S.E., Wright, W.S., and MacDougald, O.A. (2002). Wnt signaling protects 3T3-L1 preadipocytes from apoptosis through induction of insulin-like growth factors. J Biol Chem 277, 38239-38244. Lukoyanova, N., VanLoock, M.S., Orlova, A., Galkin, V.E., Wang, K., and Egelman, E.H. (2002). Each actin subunit has three nebulin binding sites: implications for steric blocking. Curr Biol 12, 383-388. Lyons, I., Parsons, L.M., Hartley, L., Li, R., Andrews, J.E., Robb, L., and Harvey, R.P. (1995). Myogenic and morphogenetic defects in the heart tubes of murine embryos lacking the homeo box gene Nkx2-5. Genes Dev 9, 1654-1666. Ma, X., Takeda, K., Singh, A., Yu, Z.X., Zerfas, P., Blount, A., Liu, C., Towbin, J.A., Schneider, M.D., Adelstein, R.S., et al. (2009). Conditional ablation of nonmuscle myosin II-B delineates heart defects in adult mice. Circ Res 105, 1102-1109. Maass, K., Shibayama, J., Chase, S.E., Willecke, K., and Delmar, M. (2007). C-terminal truncation of connexin43 changes number, size, and localization of cardiac gap junction plaques. Circ Res 101, 1283-1291. Maier, S.K., Westenbroek, R.E., McCormick, K.A., Curtis, R., Scheuer, T., and Catterall, W.A. (2004). Distinct subcellular localization of different sodium channel alpha and beta subunits in single ventricular myocytes from mouse heart. Circulation 109, 1421-1427. 236 Maier, S.K., Westenbroek, R.E., Schenkman, K.A., Feigl, E.O., Scheuer, T., and Catterall, W.A. (2002). An unexpected role for brain-type sodium channels in coupling of cell surface depolarization to contraction in the heart. Proc Natl Acad Sci U S A 99, 4073-4078. Malekar, P., Hagenmueller, M., Anyanwu, A., Buss, S., Streit, M.R., Weiss, C.S., Wolf, D., Riffel, J., Bauer, A., Katus, H.A., et al. (2010). Wnt signaling is critical for maladaptive cardiac hypertrophy and accelerates myocardial remodeling. Hypertension 55, 939-945. Malhotra, J.D., Thyagarajan, V., Chen, C., and Isom, L.L. (2004). Tyrosinephosphorylated and nonphosphorylated sodium channel beta1 subunits are differentially localized in cardiac myocytes. J Biol Chem 279, 40748-40754. Manisastry, S.M., Zaal, K.J., and Horowits, R. (2009). Myofibril assembly visualized by imaging N-RAP, alpha-actinin, and actin in living cardiomyocytes. Exp Cell Res 315, 2126-2139. Markwald, R.R., Norris, R.A., Moreno-Rodriguez, R., and Levine, R.A. (2010). Developmental basis of adult cardiovascular diseases: valvular heart diseases. Ann N Y Acad Sci 1188, 177-183. Maron, B.J., and Ferrans, V.J. (1973). Significance of multiple intercalated discs in hypertrophied human myocardium. The American journal of pathology 73, 81-96. Marvin, M.J., di Rocco, G., Gardiner, A., Bush, S.M., and Lassar, A.B. (2001). Inhibition of Wnt activity induces heart formation from posterior mesoderm. Genes & Dev 15, 316-327. Masuelli, L., Bei, R., Sacchetti, P., Scappaticci, I., Francalanci, P., Albonici, L., Coletti, A., Palumbo, C., Minieri, M., Fiaccavento, R., et al. (2003). β-catenin accumulates in intercalated disks of hypertrophic cardiomyopathic hearts. Cardiovasc Res 60, 376-387. McCalmon, S.A., Desjardins, D.M., Ahmad, S., Davidoff, K.S., Snyder, C.M., Sato, K., Ohashi, K., Kielbasa, O.M., Mathew, M., Ewen, E.P., et al. (2010). Modulation of angiotensin II-mediated cardiac remodeling by the MEF2A target gene Xirp2. Circ Res 106, 952-960. McKoy, G., Protonotarios, N., Crosby, A., Tsatsopoulou, A., Anastasakis, A., Coonar, A., Norman, M., Baboonian, C., Jeffery, S., and McKenna, W.J. (2000). Identification of a deletion in plakoglobin in arrhythmogenic right ventricular cardiomyopathy with palmoplantar keratoderma and woolly hair (Naxos disease). Lancet 355, 2119-2124. McLachlan, R.W., and Yap, A.S. (2007). Not so simple: the complexity of phosphotyrosine signaling at cadherin adhesive contacts. J Mol Med 85, 545-554. Meadows, L.S., and Isom, L.L. (2005). Sodium channels as macromolecular complexes: implications for inherited arrhythmia syndromes. Cardiovasc Res 67, 448-458. Meng, W., Mushika, Y., Ichii, T., and Takeichi, M. (2008). Anchorage of microtubule minus ends to adherens junctions regulates epithelial cell-cell contacts. Cell 135, 948-959. 237 Mertens, C., Kuhn, C., and Franke, W.W. (1996). Plakophilins 2a and 2b: constitutive proteins of dual location in the karyoplasm and the desmosomal plaque. J Cell Biol 135, 1009-1025. Mohler, P.J., Rivolta, I., Napolitano, C., LeMaillet, G., Lambert, S., Priori, S.G., and Bennett, V. (2004). Nav1.5 E1053K mutation causing Brugada syndrome blocks binding to ankyrin-G and expression of Nav1.5 on the surface of cardiomyocytes. Proc Natl Acad Sci U S A 101, 17533-17538. Mohler, P.J., Splawski, I., Napolitano, C., Bottelli, G., Sharpe, L., Timothy, K., Priori, S.G., Keating, M.T., and Bennett, V. (2004). A cardiac arrhythmia syndrome caused by loss of ankyrin-B function. Proc Natl Acad Sci U S A 101, 9137-9142. Molenaar, M., van de Wetering, M., Oosterwegel, M., Peterson-Maduro, J., Godsave, S., Korinek, V., Roose, J., Destree, O., and Clevers, H. (1996). XTcf-3 transcription factor mediates β-catenin-induced axis formation in Xenopus embryos. Cell 86, 391-399. Naito, A.T., Shiojima, I., Akazawa, H., Hidaka, K., Morisaki, T., Kikuchi, A., and Komuro, I. (2006). Developmental stage-specific biphasic roles of Wnt/betacatenin signaling in cardiomyogenesis and hematopoiesis. Proc Natl Acad Sci U S A 103, 19812-19817. Nelson, W.J., and Nusse, R. (2004). Convergence of Wnt, β-catenin, and cadherin pathways. Science 303, 1483-1487. Nerbonne, J.M., and Kass, R.S. (2005). Molecular physiology of cardiac repolarization. Physiol Rev 85, 1205-1253. Noorman, M., van der Heyden, M.A., van Veen, T.A., Cox, M.G., Hauer, R.N., de Bakker, J.M., and van Rijen, H.V. (2009). Cardiac cell-cell junctions in health and disease: Electrical versus mechanical coupling. J Mol Cell Cardiol 47, 23-31. Noren, N.K., Liu, B.P., Burridge, K., and Kreft, B. (2000). p120 catenin regulates the actin cytoskeleton via Rho family GTPases. J Cell Biol 150, 567-580. Noren, N.K., Niessen, C.M., Gumbiner, B.M., and Burridge, K. (2001). Cadherin engagement regulates Rho family GTPases. J Biol Chem 276, 33305-33308. Novy, R.E., Sellers, J.R., Liu, L.-F., and Lin, J.J.-C. (1993). In vitro functional characterization of bacterially expressed human fibroblast tropomyosin isoforms and their chimeric mutants. Cell Motil Cytoskel 26, 248-261. Olson, E.N. (2006). Gene regulatory networks in the evolution and development of the heart. Science 313, 1922-1927. Ong, L.-L., Kim, N., Mima, T., Cohen-Gould, L., and Mikawa, T. (1998). Trabecular myocytes of the embryonic heart require N-cadherin for migratory unit identity. Developmental Biology 193, 1-9. Otten, J., van der Ven, P.F., Vakeel, P., Eulitz, S., Kirfel, G., Brandau, O., Boesl, M., Schrickel, J.W., Linhart, M., Hayess, K., et al. (2010). Complete loss of murine Xin results in a mild cardiac phenotype with altered distribution of intercalated discs. Cardiovasc Res 85, 739-750. 238 Oxford, E.M., Musa, H., Maass, K., Coombs, W., Taffet, S.M., and Delmar, M. (2007). Connexin43 remodeling caused by inhibition of plakophilin-2 expression in cardiac cells. Circ Res 101, 703-711. Pacholsky, D., Vakeel, P., Himmel, M., Lowe, T., Stradal, T., Rottner, K., Furst, D.O., and van der Ven, P.F. (2004). Xin repeats define a novel actin-binding motif. J Cell Sci 117, 5257-5268. Palatinus, J.A., and Gourdie, R.G. (2007). Xin and the art of intercalated disk maintenance. Am J Physiol Heart Circ Physiol 293, H2626-2628. Pannekoek, W.J., Kooistra, M.R., Zwartkruis, F.J., and Bos, J.L. (2009). Cell-cell junction formation: the role of Rap1 and Rap1 guanine nucleotide exchange factors. Biochim Biophys Acta 1788, 790-796. Pece, S., Chiariello, M., Murga, C., and Gutkind, J.S. (1999). Activation of the protein kinase Akt/PKB by the formation of E-cadherin-mediated cell-cell junctions. Evidence for the association of phosphatidylinositol 3-kinase with the E-cadherin adhesion complex. J Biol Chem 274, 19347-19351. Perez, T.D., Tamada, M., Sheetz, M.P., and Nelson, W.J. (2008). Immediate-early signaling induced by E-cadherin engagement and adhesion. J Biol Chem 283, 5014-5022. Perez-Moreno, M., and Fuchs, E. (2006). Catenins: keeping cells from getting their signals crossed. Dev Cell 11, 601-612. Perriard, J.C., Hirschy, A., and Ehler, E. (2003). Dilated cardiomyopathy: a disease of the intercalated disc? Trends Cardiovasc Med 13, 30-38. Peters, N.S., Severs, N.J., Rothery, S.M., Lincoln, C., Yacoub, M.H., and Green, C.R. (1994). Spatiotemporal relation between gap junctions and fascia adherens junctions during postnatal development of human ventricular myocardium. Circulation 90, 713-725. Pieperhoff, S., and Franke, W.W. (2007). The area composita of adhering junctions connecting heart muscle cells of vertebrates - IV: coalescence and amalgamation of desmosomal and adhaerens junction components - late processes in mammalian heart development. Eur J Cell Biol 86, 377-391. Pieperhoff, S., and Franke, W.W. (2008). The area composita of adhering junctions connecting heart muscle cells of vertebrates. VI. Different precursor structures in non-mammalian species. Eur J Cell Biol 87, 413-430. Pieperhoff, S., Schumacher, H., and Franke, W.W. (2008). The area composita of adhering junctions connecting heart muscle cells of vertebrates. V. The importance of plakophilin-2 demonstrated by small interference RNA-mediated knockdown in cultured rat cardiomyocytes. Eur J Cell Biol 87, 399-411. Pilichou, K., Remme, C.A., Basso, C., Campian, M.E., Rizzo, S., Barnett, P., Scicluna, B.P., Bauce, B., van den Hoff, M.J., de Bakker, J.M., et al. (2009). Myocyte necrosis underlies progressive myocardial dystrophy in mouse dsg2-related arrhythmogenic right ventricular cardiomyopathy. J Exp Med 206, 1787-1802. 239 Pokutta, S., and Weis, W.I. (2000). Structure of the dimerization and α-catenin-binding region of α-catenin. Mol Cell 5, 533-543. Pokutta, S., and Weis, W.I. (2002). The cytoplasmic face of cell contact sites. Curr Opin Struc Biol 23, 255-262. Pollard, T.D., Blanchoin, L., and Mullins, R.D. (2000). Molecular mechanisms controlling actin filament dynamics in nonmuscle cells. Annu Rev Biophys Biomol Struct 29, 545-576. Porrello, E.R., Mahmoud, A.I., Simpson, E., Hill, J.A., Richardson, J.A., Olson, E.N., and Sadek, H.A. (2011). Transient regenerative potential of the neonatal mouse heart. Science 331, 1078-1080. Qu, J., Zhou, J., Yi, X.P., Dong, B., Zheng, H., Miller, L.M., Wang, X., Schneider, M.D., and Li, F. (2007). Cardiac-specific haploinsufficiency of beta-catenin attenuates cardiac hypertrophy but enhances fetal gene expression in response to aortic constriction. J Mol Cell Cardiol 43, 319-326. Rambaut, A. (1995). SE-AL sequence alignment program. Version 1.D1 (Department of Zoology, University of Oxford, Oxford, England). Raptis, L., Arulanandam, R., Vultur, A., Geletu, M., Chevalier, S., and Feracci, H. (2009). Beyond structure, to survival: activation of Stat3 by cadherin engagement. Biochem Cell Biol 87, 835-843. Reynolds, A.B., and Roczniak-Ferguson, A. (2004). Emerging roles for p120-catenin in cell adhesion and cancer. Oncogene 23, 7947-7956. Rockman, H.A., Knowlton, K.U., J. Ross, J., and Chien, K.R. (1993). In vivo murine cardiac hypertrophy. Circulation 87[suppl VII], VII14-VII21. Rockman, H.A., Ross, R.S., Harris, A.N., Knowlton, K.U., Steinhelper, M.E., Field, L.J., Ross, J., Jr., and Chien, K.R. (1991). Segregation of atrial-specific and inducible expression of an atrial natriuretic factor transgene in an in vivo murine model of cardiac hypertrophy. Proc Natl Acad Sci U S A 88, 8277-8281. Rohr, S. (2007). Molecular crosstalk between mechanical and electrical junctions at the intercalated disc. Circ Res 101, 637-639. Ross, S.E., Hemati, N., Longo, K.A., Bennett, C.N., Lucas, P.C., Erickson, R.L., and MacDougald, O.A. (2000). Inhibition of adipogenesis by Wnt signaling. Science 289, 950-953. Rubinfeld, B., Souza, B., Albert, I., Muller, O., Chamberlain, S.H., Masiarz, F.R., Munemitsu, S., and Polakis, P. (1993). Association of the APC gene product with β-catenin. Science 262, 1731-1734. Ruiz, P., Brinkmann, V., Ledermann, B., Behrend, M., Grund, C., Thalhammer, C., Vogel, F., Birchmeier, C., Gunthert, U., Franke, W.W., et al. (1996). Targeted mutation of plakoglobin in mice reveals essential functions of desmosomes in the embryonic heart. J Cell Biol 135, 215-225. 240 Sadot, E., Simcha, I., Shtutman, M., Ben-Ze'ev, A., and Geiger, B. (1998). Inhibition of β-catenin-mediated transactivation by cadherin derivatives. Proc Natl Acad Sci USA 95, 15339-15344. Sato, P.Y., Coombs, W., Lin, X., Nekrasova, O., Green, K.J., Isom, L.L., Taffet, S.M., and Delmar, M. (2011). Interactions between ankyrin-G, Plakophilin-2, and Connexin43 at the cardiac intercalated disc. Circ Res 109, 193-201. Sato, P.Y., Musa, H., Coombs, W., Guerrero-Serna, G., Patino, G.A., Taffet, S.M., Isom, L.L., and Delmar, M. (2009). Loss of plakophilin-2 expression leads to decreased sodium current and slower conduction velocity in cultured cardiac myocytes. Circ Res 105, 523-526. Satoh, M., Ogita, H., Takeshita, K., Mukai, Y., Kwiatkowski, D.J., and Liao, J.K. (2006). Requirement of Rac1 in the development of cardiac hypertrophy. Proc Natl Acad Sci U S A 103, 7432-7437. Severs, N.J. (1990). The cardiac gap junction and intercalated disc. Int J Cardiol 26, 137173. Severs, N.J., Bruce, A.F., Dupont, E., and Rothery, S. (2008). Remodelling of gap junctions and connexin expression in diseased myocardium. Cardiovasc Res 80, 9-19. Shaw, R.M., Fay, A.J., Puthenveedu, M.A., von Zastrow, M., Jan, Y.N., and Jan, L.Y. (2007). Microtubule plus-end-tracking proteins target gap junctions directly from the cell interior to adherens junctions. Cell 128, 547-560. Sheikh, F., Chen, Y., Liang, X., Hirschy, A., Stenbit, A.E., Gu, Y., Dalton, N.D., Yajima, T., Lu, Y., Knowlton, K.U., et al. (2006). alpha-E-catenin inactivation disrupts the cardiomyocyte adherens junction, resulting in cardiomyopathy and susceptibility to wall rupture. Circulation 114, 1046-1055. Sheikh, F., Ross, R.S., and Chen, J. (2009). Cell-cell connection to cardiac disease. Trends Cardiovasc Med 19, 182-190. Shih, I.-M., Yu, J., He, T.-C., Vogelstein, B., and Kinzler, K.W. (2000). The β-catenin binding domain of adenomatous polyposis coli is sufficient for tumor suppression. Cancer Res 60, 1671-1676. Sinn, H.W., Balsamo, J., Lilien, J., and Lin, J.J. (2002). Localization of the novel Xin protein to the adherens junction complex in cardiac and skeletal muscle during development. Dev Dyn 225, 1-13. Smith, J.H., Green, C.R., Peters, N.S., Rothery, S., and Severs, N.J. (1991). Altered patterns of gap junction distribution in ischemic heart disease. An immunohistochemical study of human myocardium using laser scanning confocal microscopy. Am J Pathol 139, 801-821. Smutny, M., and Yap, A.S. (2010). Neighborly relations: cadherins and mechanotransduction. J Cell Biol 189, 1075-1077. Soll, D.R. (1995). The use of computers in understanding how animal cells crawl. International Review of Cytology 8, 439-454. 241 Soll, D.R., and Voss, E. (1998). Two and three-dimensional computer systems for analyzing how cells crawl. In Motion analysis of living cells, D. Soll, and D. Wessels, eds. (New York, Wiley-Liss), pp. 25-52. Somekawa, S., Fukuhara, S., Nakaoka, Y., Fujita, H., Saito, Y., and Mochizuki, N. (2005). Enhanced functional gap junction neoformation by protein kinase Adependent and Epac-dependent signals downstream of cAMP in cardiac myocytes. Circ Res 97, 655-662. Sonnenberg, A., and Liem, R.K. (2007). Plakins in development and disease. Exp Cell Res 313, 2189-2203. Srivastava, D. (2006). Making or breaking the heart: from lineage determination to morphogenesis. Cell 126, 1037-1048. Stamatakis, A., Ludwig, T., and Meier, H. (2005). RAxML-III: a fast program for maximum likelihood-based inference of large phylogenetic trees. Bioinformatics 21, 456-463. Stevenson, B.R., Siliciano, J.D., Mooseker, M.S., and Goodenough, D.A. (1986). Identification of ZO-1: a high molecular weight polypeptide associated with the tight junction (zonula occludens) in a variety of epithelia. J Cell Biol 103, 755766. Stossel, T.S., Condeelis, J., Cooley, L., Hartwig, J.H., Noegel, A., Schleicher, M., and Shapiro, S.S. (2001). Filamins as integrators of cell mechanics and signalling. Nat Rev Mol Cell Biol 2, 138-145. Su, L.-K., Vogelstein, B., and Kinzler, K.W. (1993). Association of the APC tumor suppressor protein with catenins. Science 262, 1734-1737. Sussman, M.A., Welch, S., Walker, A., Klevitsky, R., Hewett, T.E., Price, R.L., Schaefer, E., and Yager, K. (2000). Altered focal adhesion regulation correlates with cardiomyopathy in mice expressing constitutively active rac1. J Clin Invest 105, 875-886. Thompson, J.D., Higgins, D.G., and Gibson, T.J. (1994). CLUSTAL W: improving the sensitivity of progressive multiple sequence alignments through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Research 22, 4673-4680. Toyofuku, T., Yabuki, M., Otsu, K., Kuzuya, T., Hori, M., and Tada, M. (1998). Direct association of the gap junction protein connexin-43 with ZO-1 in cardiac myocytes. J Biol Chem 273, 12725-12731. Tzahor, E., and Lassar, A.B. (2001). Wnt signals from the neural tube block ectopic cardiogenesis. Genes & Dev 15, 255-260. Ueno, S., Weidinger, G., Osugi, T., Kohn, A.D., Golob, J.L., Pabon, L., Reinecke, H., Moon, R.T., and Murry, C.E. (2007). Biphasic role for Wnt/beta-catenin signaling in cardiac specification in zebrafish and embryonic stem cells. Proc Natl Acad Sci U S A 104, 9685-9690. 242 van de Schans, V.A., van den Borne, S.W., Strzelecka, A.E., Janssen, B.J., van der Velden, J.L., Langen, R.C., Wynshaw-Boris, A., Smits, J.F., and Blankesteijn, W.M. (2007). Interruption of Wnt signaling attenuates the onset of pressure overload-induced cardiac hypertrophy. Hypertension 49, 473-480. van der Flier, A., Kuikman, I., Kramer, D., Geerts, D., Kreft, M., Takafuta, T., Shapiro, S.S., and Sonnenberg, A. (2002). Different splice variants of filamin-B affect myogenesis, subcellular distribution, and determine binding to integrin β subunits. J Cell Biol 136, 361-376. van der Heyden, M.A., Rook, M.B., Hermans, M.M., Rijksen, G., Boonstra, J., Defize, L.H., and Destree, O.H. (1998). Identification of connexin43 as a functional target for Wnt signalling. J Cell Sci 111 ( Pt 12), 1741-1749. van der Ven, P.F.M., Ehler, E., Vakeel, P., Eulitz, S., Schenk, J.A., Milting, H., Micheel, B., and Furst, D.O. (2006). Unusual splicing events result in distinct Xin isoforms that associate differentially with filamin c and Mens/VASP. Exp Cell Res 312, 2154-2167. van Tintelen, J.P., Entius, M.M., Bhuiyan, Z.A., Jongbloed, R., Wiesfeld, A.C., Wilde, A.A., van der Smagt, J., Boven, L.G., Mannens, M.M., van Langen, I.M., et al. (2006). Plakophilin-2 mutations are the major determinant of familial arrhythmogenic right ventricular dysplasia/cardiomyopathy. Circulation 113, 1650-1658. Wang, D.Z., Hu, X., Lin, J.L., Kitten, G.T., Solursh, M., and Lin, J.J. (1996). Differential displaying of mRNAs from the atrioventricular region of developing chicken hearts at stages 15 and 21. Front Biosci 1, a1-15. Wang, D.-Z., Reiter, R.S., Lin, J.L.-C., Wang, Q., Williams, H.S., Krob, S.L., Schultheiss, T.M., Evans, S., and Lin, J.J.-C. (1999). Requirement of a novel gene, Xin, in cardiac morphogenesis. Development 126, 1281-1294. Wang, Q., Lin, J.L., Reinking, B.E., Feng, H.Z., Chan, F.C., Lin, C.I., Jin, J.P., Gustafson-Wagner, E.A., Scholz, T.D., Yang, B., et al. (2010). Essential roles of an intercalated disc protein, mXinβ, in postnatal heart growth and survival. Circ Res 16, 1468-1478. Wang, X., and Gerdes, A.M. (1999). Chronic pressure overload cardiac hypertrophy and failure in guinea pigs: III. intercalated disc remodeling. J Mol Cell Cardiol 31, 333-343. Warren, K.S., and Lin, J.J.-C. (1993). Forced expression and assembly of rat cardiac troponin T isoforms in cultured muscle and nonmuscle cells. Journal of Muscle Research and Cell Motility 14, 619-632. Watanabe, T., Sato, K., and Kaibuchi, K. (2009). Cadherin-mediated intercellular adhesion and signaling cascades involving small GTPases. Cold Spring Harb Perspect Biol 1, a003020. Wei, C.J., Francis, R., Xu, X., and Lo, C.W. (2005). Connexin43 associated with an Ncadherin-containing multiprotein complex is required for gap junction formation in NIH3T3 cells. J Biol Chem 280, 19925-19936. 243 Westfall, M.V., and Metzger, J.M. (2001). Troponin I isoforms and chimeras: tuning the molecular switch of cardiac contraction. News Physiol Sci 16, 278-281. Wheelock, M.J., and Johnson, K.R. (2003). Cadherin-mediated cellular signaling. Curr Opin Cell Biol 15, 509-514. Wu, J.C., Tsai, R.Y., and Chung, T.H. (2003). Role of catenins in the development of gap junctions in rat cardiomyocytes. J Cell Biochem 88, 823-835. Xing, Y., Clements, W.K., Kimelman, D., and Xu, W. (2003). Crystal structure of a βcatenin/axin complex suggests a mechanism for the β-catenin destruction complex. Genes & Dev 17, 2753-2764. Xu, Y., Guo, D.F., Davidson, M., Inagami, T., and Carpenter, G. (1997). Interaction of the adaptor protein Shc and the adhesion molecule cadherin. J Biol Chem 272, 13463-13466. Yamada, K.A., Rogers, J.G., Sundset, R., Steinberg, T.H., and Saffitz, J.E. (2003). Upregulation of connexin45 in heart failure. J Cardiovasc Electrophysiol 14, 12051212. Yamada, S., and Nelson, W.J. (2007). Localized zones of Rho and Rac activities drive initiation and expansion of epithelial cell-cell adhesion. J Cell Biol 178, 517-527. Yamada, S., Pokutta, S., Drees, F., Weis, W.I., and Nelson, W.J. (2005). Deconstructing the cadherin-catenin-actin complex. Cell 123, 889-901. Yamane, M., Matsuda, T., Ito, T., Fujio, Y., Takahashi, K., and Azuma, J. (2007). Rac1 activity is required for cardiac myocyte alignment in response to mechanical stress. Biochem Biophys Res Commun 353, 1023-1027. Yang, Z., Bowles, N.E., Scherer, S.E., Taylor, M.D., Kearney, D.L., Ge, S., Nadvoretskiy, V.V., DeFreitas, G., Carabello, B., Brandon, L.I., et al. (2006). Desmosomal dysfunction due to mutations in desmoplakin causes arrhythmogenic right ventricular dysplasia/cardiomyopathy. Circ Res 99, 646-655. Yonemura, S., Wada, Y., Watanabe, T., Nagafuchi, A., and Shibata, M. (2010). alphaCatenin as a tension transducer that induces adherens junction development. Nat Cell Biol 12, 533-542. Yoshida, M., Sho, E., Nanjo, H., Takahashi, M., Kobayashi, M., Kawamura, K., Honma, M., Komatsu, M., Sugita, A., Yamauchi, M., et al. (2010). Weaving hypothesis of cardiomyocyte sarcomeres: discovery of periodic broadening and narrowing of intercalated disk during volume-load change. Am J Pathol 176, 660-678. Zemljic-Harpf, A.E., Miller, J.C., Henderson, S.A., Wright, A.T., Manso, A.M., Elsherif, L., Dalton, N.D., Thor, A.K., Perkins, G.A., McCulloch, A.D., et al. (2007). Cardiac-myocyte-specific excision of the vinculin gene disrupts cellular junctions, causing sudden death or dilated cardiomyopathy. Mol Cell Biol 27, 7522-7537. Zhang, J.Q., Elzey, B., Williams, G., Lu, S., Law, D.J., and Horowits, R. (2001). Ultrastructural and biochemical localization of N-RAP at the interface between myofibrils and intercalated disks in the mouse heart. Biochemistry 40, 1489814906. 244 Zhou, J., Qu, J., Yi, X.P., Graber, K., Huber, L., Wang, X., Gerdes, A.M., and Li, F. (2007). Upregulation of gamma-catenin compensates for the loss of beta-catenin in adult cardiomyocytes. Am J Physiol Heart Circ Physiol 292, H270-276. Zhou, Y.Q., Foster, F.S., Parkes, R., and Adamson, S.L. (2003). Developmental changes in left and right ventricular diastolic filling patterns in mice. Am J Physiol Heart Circ Physiol 285, H1563-1575. Zhurinsky, J., Shtutman, M., and Ben-Ze'ev, A. (2000). Plakoglobin and beta-catenin: protein interactions, regulation and biological roles. J Cell Sci 113 ( Pt 18), 31273139.