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Intercellular junction disassembly : assigning a function to apical tubulobulbar complexes in testis Young, J'Nelle Sarah 2013

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INTERCELLULAR JUNCTION DISASSEMBLY: ASSIGNING A FUNCTION TO APICAL TUBULOBULBAR COMPLEXES IN TESTIS  by  JʼNelle Sarah Young  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY  in  The Faculty of Graduate Studies  (Cell and Developmental Biology)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  April 2013  © JʼNelle Sarah Young, 2013  Abstract 	
   The focus of this thesis is the characterization of apical tubulobulbar complexes in mammalian testis. These double-membrane, actin-based structures form at sites of attachment between germ cells and Sertoli cells. The location, timing and morphology of these complexes have inspired several proposed functions. It has been proposed that tubulobulbar complexes serve as an anchor to prevent early release of spermatids from the epithelium or that they are a device for elimination of excess cytoplasm. I hypothesize that tubulobulbar complexes are subcellular machines responsible for the internalization of intact intercellular junctions thereby contributing to the process of spermatid release from the seminiferous epithelium. A descriptive approach is taken to determine if key components that are present at similar structures in other systems also are present at tubulobulbar complexes, to determine if integral junction molecules are present at tubulobulbar complexes and to determine the fate of internalized junction material.  A  functional approach is taken to deplete the expression of an actin-based protein that is localized to tubulobulbar complexes to test the prediction that the structures are involved with spermatid release. Dendritic actin components were localized around the cuff of tubulobulbar complexes and clathrin was localized to coated pits at the ends of the structures. Based on these data, a model of tubulobulbar complex formation was proposed that incorporates clathrin-mediated endocytosis and dendritic actin assembly. 	
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    Junction molecules known to be present at ectoplasmic specializations were found in vesicles at the ends of tubulobulbar complexes that label positively for early endosome markers. Interestingly, junction protein nectin 2 was colocalized with recycling marker Rab11 at newly forming junctions deeper in the epithelium. This suggests that recycling of junctional proteins may be occurring in Sertoli cells. Finally, depletion of cortactin, a key protein at tubulobulbar complexes, resulted in a short phenotype – an indication that the structures were not able to acquire or maintain their normal length after treatment. Significantly, delay in spermatid release was detected. The data presented here supports the junction internalization hypothesis and introduces a new paradigm for junction internalization generally in cells and links the mechanism to a biologically significant event – sperm release.  	
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    Preface  Chapter 2: Cortactin (CTTN), N-WASP (WASL), and Clathrin (CLTC) Are Present at Podosome-Like Tubulobulbar Complexes in the Rat Testis Accepted for publication in Biology of Reproduction, September 5, 2008. JʼNelle S. Young, Julian A. Guttman, Kuljeet S. Vaid, Hasmik Shahinian, and A. Wayne Vogl Author contributions: JʼNelle Young was involved in execution of experiments, data analysis, manuscript writing and editing. Julian Guttman and Kuljeet Vaid contributed to experimental design. Hasmik Shahinian was involved in execution of experiments for the majority of Clathrin studies. A. Wayne Vogl was involved in experimental design, execution of each of the Cortactin, N-WASP and Clathrin studies, data analysis, manuscript writing and editing.  Chapter 3: Focal Adhesion Proteins Zyxin and Vinculin Are Co-distributed at Tubulobulbar Complexes in Rat Testis. Accepted for publication in Spermatogenesis, January 17, 2012. JʼNelle S. Young and A. Wayne Vogl Author contributions: JʼNelle Young was involved in experimental design, execution of experiments, data analysis, writing and editing of manuscript. A. Wayne Vogl was involved experimental design, execution of experiments, in data analysis, and editing manuscript. 	
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    Chapter 4: Tubulobulbar Complexes are Intercellular Podosome-Like Structures That Internalize Intact Intercellular Junctions During Epithelial Remodeling Events in Testis. Accepted for publication in Biology of Reproduction, September 9, 2008. JʼNelle S. Young, Julian A. Guttman, Kuljeet S. Vaid, and A. Wayne Vogl Author contributions: JʼNelle Young was involved in execution of experiments, data analysis, writing and editing of manuscript. Kuljeet Vaid and Julian Guttman contributed to experimental design. A. Wayne Vogl was involved in experimental design, execution of experiments, analysis of data and writing and editing of the manuscript.  Chapter 5: Internalization of Adhesion Junction Proteins and Their Association with Recycling Endosome Marker Proteins in Rat Seminiferous Epithelium. Accepted for publication in Reproduction, December 12, 2011 JʼNelle S. Young, Yoshimi Takai, Katarina L. Kojic and A. Wayne Vogl Author contributions: JʼNelle Young was involved in experimental design, execution of experiments, data analysis, writing and editing of the manuscript. Yoshimi Takai supplied nectin2 antibody, edited manuscript and contributed to experimental design. Katarina Kojic was involved with execution of β1 integrin studies and data  	
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    analysis. A. Wayne Vogl was involved with experimental design, execution of experiments, data analysis, writing and editing of manuscript.  Chapter 6: Cortactin Depletion Results in Short Tubulobulbar Complexes and Spermiation Failure in Rat Testes. Accepted for publication in Biology Open, July 16, 2012 JʼNelle S. Young, Marc de Asis, Julian Guttman and A. Wayne Vogl Author contributions: JʼNelle Young was involved in experimental design, execution of experiments, data analysis, and writing and editing of manuscript. Marc de Asis assisted with animal surgeries and edited manuscript. Julian Guttman contributed to experimental design, A. Wayne Vogl was involved with experimental design, execution of experiments, data analysis, and writing and editing of manuscript.  	
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    Table of Contents  Abstract................................................................................................................ ii Preface ................................................................................................................ iv Table of Contents.............................................................................................. vii List of Figures .................................................................................................. xiii Acknowledgements ....................................................................................... xviii Dedication......................................................................................................... xix Chapter 1 Introduction .................................................................................... 1 1.1 Organization of the mammalian seminiferous epithelium and spermatogenesis............................................................................................. 1 1.2 Arrangement and functional significance of intercellular junctions in the seminiferous epithelium .......................................................................... 2 1.3 Intercellular junction complexes ............................................................. 2 1.3.1 Tight junctions ....................................................................................... 3 1.3.2 Adherens junctions ............................................................................... 3 1.3.3 Gap junctions ........................................................................................ 4 1.3.4 Desmosomes ........................................................................................ 6 1.4 Intercellular junctions in the seminiferous epithelium.......................... 6  	
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    1.4.1 Ectoplasmic specializations .................................................................. 7 1.5 Junction turnover in epithelia .................................................................. 9 1.5.1 The endosomal network........................................................................ 9 1.6 Tubulobulbar complexes........................................................................ 11 1.6.1 Dendritic actin assembly ..................................................................... 12 1.6.2 Basal tubulobulbar complexes ............................................................ 13 1.6.3 Apical tubulobulbar complexes ........................................................... 14 1.7 Morphologically similar structures........................................................ 15 1.7.1 Clathrin-mediated endocytosis machinery .......................................... 16 1.7.2 Podosomes formed by osteoclasts ..................................................... 17 1.7.3 Reconstituted membrane tubules ....................................................... 18 1.8 Proposed functions of tubulobulbar complexes.................................. 19 1.9 Working hypothesis and chapter aims ................................................. 20 1.10 Introduction figures............................................................................... 26 Chapter 2 – Cortactin (CTTN), n-wasp (WASL) and clathrin (CLTC) are present at podosome-like tubulobulbar complexes in rat testis ................. 30 2.1 Brief synopsis.......................................................................................... 30 2.2 Results...................................................................................................... 32 2.2.1 Tubulobulbar complexes contain n-wasp and cortactin ...................... 32 2.2.2 Tubulobulbar complexes contain cortactin.......................................... 32 2.2.3 Tubulobulbar complexes contain n-wasp............................................ 33 2.2.4 Clathrin is localized to the ends of tubulobulbar complexes ............... 34  	
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    2.3 Discussion ............................................................................................... 35 2.3 Chapter 1 figures ..................................................................................... 39 Chapter 3 – Focal adhesion proteins zyxin and vinculin are co-distributed at tubulobulbar complexes in rat testis .......................................................... 49 3.1 Brief synopsis.......................................................................................... 49 3.2 Results...................................................................................................... 51 3.2.1 The zyxin antibody labels ectoplasmic specializations and tubulobulbar complexes.................................................................................................... 51 3.2.2 Zyxin is present with vinculin at tubulobulbar complexes ................... 52 3.2.3 In testis, the zyxin antibody is specific for zyxin splice variant HED-2.52 3.3 Discussion ............................................................................................... 53 3.4 Chapter 3 figures ..................................................................................... 56 Chapter 4 – Tubulobulbar complexes are intercellular podosome-like structures that internalize intact intercellular junctions during epithelial remodeling events in testis. ............................................................................ 60 4.1 Brief synopsis.......................................................................................... 60 4.2 Results...................................................................................................... 62 4.2.1 Nectin 2 ............................................................................................... 62 4.2.2 Nectin 3 ............................................................................................... 63 4.2.3 α6 integrin ........................................................................................... 64 4.2.4 N-Cadherin is concentrated at desmosome junctions and not at ectoplasmic specializations or tubulobulbulbar complexes.......................... 64  	
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    4.3 Discussion ............................................................................................... 66 4.4 Chapter 3 figures ..................................................................................... 72 Chapter 5 – Internalization of adhesion junction proteins and their association with recycling endosome marker proteins in rat seminiferous epithelium. ......................................................................................................... 86 5.1 Brief synopsis.......................................................................................... 86 5.2 Results...................................................................................................... 87 5.2.1 Nectin 2 and the β1 integrin subunit are concentrated at the ends of apical tubulobulbar complexes..................................................................... 87 5.2.2 The early endosomal marker rab5 is distinctly localized at the ends of tubulobulbar complexes. .............................................................................. 89 5.2.3 The ʻlong-loopʼ recycling marker rab11 is localized to newly forming junctions associated with early spermatids.................................................. 89 5.2.4 Rab11 is co-distributed with nectin 2 at newly forming junctions ........ 90 5.3 Discussion ............................................................................................... 91 5.4 Chapter 5 figures ..................................................................................... 96 Chapter 6 – Cortactin depletion results in short tubulobulbar complexes and spermiation failure in rat testes. ............................................................ 108 6.1 Brief synopsis........................................................................................ 108 6.2 Results.................................................................................................... 109 6.2.1 Normal arrangement and structure of tubulobulbar complexes in rat 109 6.2.2 Cortactin knockdown by intratesticular siRNA .................................. 110  	
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    6.2.3 Tubulobulbar complexes are shorter in cortactin knockdown testes. 110 6.2.4 Spermatids retained in the epithelium............................................... 112 6.2.5 SiRNA reagents enter sertoli cells and result in cortactin knockdown in isolated seminiferous epithelia ................................................................... 113 6.3 Discussion ............................................................................................. 114 6.4 Chapter 5 figures ................................................................................... 119 Chapter 7 – Materials and methods .............................................................. 126 7.1 Animals................................................................................................... 126 7.2 Reagents ................................................................................................ 126 7.3 Primary antibodies ................................................................................ 127 7.4 Immunofluorescence ............................................................................ 128 7.4.1 Tissue preparation: ........................................................................... 128 7.4.2 Frozen sections:................................................................................ 128 7.4.3 Fragmented material: ........................................................................ 129 7.4.4 Immunostaining:................................................................................ 129 7.4.5 Antigen retrieval ................................................................................ 130 7.5 Western blotting .................................................................................... 131 7.5.1 Whole testis lysate ............................................................................ 131 7.5.2 Seminiferous epithelial lysate............................................................ 131 7.6 Light and standard electron microscopy............................................ 133 7.7 Electron microscopy ............................................................................. 134 7.8 SiRNA ..................................................................................................... 134  	
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    7.9 PEI – mediated siRNA delivery ............................................................ 135 7.10 Animal surgeries ................................................................................. 136 7.10.1 Handling:......................................................................................... 136 7.10.2 Surgical preparation: ....................................................................... 136 7.10.3 Surgical procedure: ......................................................................... 137 7.10.4 Post-surgical care and processing:................................................. 137 7.10.5 Relative quantitation and statistical analysis of actin and cortactin fluorescence as an indication of tubulobulbar complex length................... 138 Chapter 8 - concluding chapter ..................................................................... 139 8.1 Overall analysis and conclusions in light of current research ......... 139 8.1.1 Structure............................................................................................ 139 8.1.2 Function ............................................................................................ 140 8.2 Conclusions based on hypothesis presented in the introduction ... 142 8.3 Comments on strengths and limitations of the thesis ...................... 146 8.4 Future directions: .................................................................................. 147 8.5 Discussion of potential applications ................................................... 151 8.6 Specific contributions........................................................................... 151 Bibliography .................................................................................................... 153 Appendix.......................................................................................................... 164  	
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    List of Figures 	
   Figure 1.1 Intercellular junctions in epithelia in general. ..................................... 26 Figure 1.2 Schematic diagram showing a section of seminiferous epithelium. .. 27 Figure 1.3 Diagram of the structure of tubulobulbar complexes in the seminiferous epithelium. .............................................................................. 28 Figure 1.4 Ultrastructural features of tubulobulbar complexes associated with Sertoli/spermatid attachment sites at the apex of rat seminiferous epithelium. ..................................................................................................................... 29 Figure 2.1 Cortactin immunofluorescence and western blot .............................. 39 Figure 2.2 Cortactin immunofluorescence stage V ............................................. 40 Figure 2.3 Western blot of rat testis and seminiferous epithelium labeled with a nwasp antibody and normal rabbit IgG (NRIgG). ........................................... 41 Figure 2.4 Paired phase and immunofluorescence micrographs of rat seminiferous epithelium labeled for n-wasp. ................................................ 42 Figure 2.5 Paired phase and immunofluorescence micrographs of epithelial fragments of rat seminiferous epithelium containing late spermatids and associated sertoli cell regions labeled for n-wasp........................................ 43 Figure 2.6 Immunofluorescent and immunoelectron microscopic demonstration of clathrin at apical tubulobulbar complexes in the rat. .................................... 44 Figure 2.7 Known components and structural features of tubulobulbar complexes together with a diagram showing the formation of the complexes. .............. 47 	
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    Figure 2.8 Model of how dendritic actin dynamics, clathrin coat proteins, and peripheral plasma membrane proteins may interact to generate tubulobulbar complexes.................................................................................................... 48 Figure 3.1 Fluorescence micrographs of rat seminiferous epithelium triple labeled for actin (phalloidin), zyxin and DNA (DAPI). ............................................... 56 Figure 3.2 Phase and immunofluorescence micrographs of a rat spermatid and attached sertoli cell regions labeled for zyxin (red), actin (green), and DNA (blue). ........................................................................................................... 57 Figure 3.3 Paired phase and immunofluorescence micrographs of rat spermatids and attached Sertoli cell regions triple labeled for zyxin, vinculin and dapi (blue). ........................................................................................................... 58 Figure 3.4 Controls for zyxin labeling of rat seminiferous epithelial fragments. .. 59 Figure 4.1 Phase and immunofluorescence micrographs of mouse spermatids and attached sertoli cell regions triple-labeled ............................................. 73 Figure 4.2 Paired phase and immunofluoresence micrographs of mouse epithelial fragments single labeled for nectin 2. .......................................................... 74 Figure 4.3 Phase and immunofluorescence micrographs of mouse spermatids and attached sertoli cell regions triple labeled ............................................. 75 Figure 4.4 Paired phase and immunofluorescence micrographs of mouse epithelial fragments containing late spermatids and adjacent sertoli cell regions single labeled for nectin 3. .............................................................. 77  	
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    Figure 4.5 Epithelial fragments of rat seminiferous epithelium triple labeled for actin, α6 integrin, and DNA (DAPI). ............................................................. 78 Figure 4.6 Controls for α6 labeling of rat seminiferous epithelial fragments. ...... 79 Figure 4.7 Paired phase and immunofluorescence micrographs of fixed frozen sections of rat testis single labeled for N-cadherin. ..................................... 80 Figure 4.8 Phase and fluorescence micrographs of rat seminiferous epithelium triple labeled for actin, n-cadherin, and DNA (DAPI).................................... 81 Figure 4.9 Immunoelectron microscopic images of rat testis sections labeled for n-cadherin. ................................................................................................... 83 Figure 4.10 Immunoelectron microscopic images of rat testis double labeled for n-cadherin and actin. ................................................................................... 84 Figure 5.1 Grouped phase and fluorescence images showing the distribution of nectin 2 and actin in rat seminiferous epithelium. ........................................ 96 Figure 5.2 Paired phase and fluorescence micrographs of late spermatids and associated Sertoli cell regions of the rat processed by the antigen retrieval protocol as indicated in the text and labeled with β1 antibodies. ................. 98 Figure 5.3 Phase and fluorescence images of late spermatids and associated sertoli lobes mechanically dissociated from perfusion fixed rat seminiferous epithelium and labeled for rab 5................................................................... 99 Figure 5.4 Paired phase and fluorescence micrographs of early spermatids labeled for actin, rab 11 and DNA. ............................................................. 102  	
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    Figure 5.5 Phase and fluorescence micrographs of spermatids and associated sertoli cell regions double labeled with antibodies to nectin 2 and rab 11. 103 Figure 5.6 Immunofluorescence micrographs of early spermatids at step 8 (a) and step 9 (b) of spermiogenesis double-labeled for nectin 2 and rab 11. 104 Figure 5.7 Shown here is a cryo-section of a seminiferous tubule containing approximately at stage XI containing step 11 spermatids.......................... 105 Figure 5.8 Cryo-sections of seminiferous tubules at stages V, VII and IX ........ 106 Figure 5.9 Model of junction turnover and recycling in the seminiferous epithelium of the rat based partly on data presented in this chapter........................... 107 Figure 6.1 Position and arrangement of tubulobulbar complexes associated with sertoli/spermatid attachment sites in rat seminiferous epithelium.............. 119 Figure 6.2 Evidence of cortactin knockdown..................................................... 120 Figure 6.3 Representative longitudinal sections of tubulobulbar complexes from two different rats. ....................................................................................... 121 Figure 6.4 Cross sections through apical sertoli cells from control and siRNA treated testes. ............................................................................................ 122 Figure 6.5 Evidence of delayed spermiation. .................................................... 123 Figure 6.6 Evidence of spermiation failure and persistence of adhesion junctions in cortactin siRNA testes. ........................................................................... 124 Figure 6.7 Cortactin knockdown occurs in the seminiferous epithelium and siRNA reagents enter Sertoli cells......................................................................... 125 Figure A.1 Sertoli cells transfected with nectin2-GFP cDNA............................. 164  	
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    Acknowledgements  First and foremost, I offer my sincere gratitude to Dr. Wayne Vogl for providing the dedication and mentorship that was necessary for me to complete this work. I continue to be inspired by his energy, enthusiasm and commitment to producing the best possible work. Wayneʼs contribution to my education and personal development will undoubtedly have an impact on the rest of my career. I would like to thank my committee members Dr. Lacey Samuels, Dr. Cal Roskelley, and Dr. Tim OʼConnor for the constructive suggestions and insight that have strengthened my thesis. I would also like to thank all of the past and present members of the Vogl lab for the great company and constant support. I offer my gratitude to the faculty, staff, and fellow students in the Life Sciences Institute who have contributed ideas and support to my project whether during a brief discussion in the hallway, at lunch or over coffee. I owe a special thanks to my family - David and Suellen and siblings, Anna, Stephanie, David and Christopher for creating and fostering an environment that values the most important things in life – love, laughter, travel and education – “top of the class!”. Finally, I would like to thank Andrew Caruthers for being my rock and believing in me – especially during the times when I had doubts in myself.  	
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    Dedication 	
   To Esther, Charlie, Anna and Sam for equipping me with the wit, kindness, grace and humor that were necessary for coping with the challenges of producing and communicating this work. To my mom and dad for their unconditional support and encouragement throughout my winding path to realizing my dream. And to Monkey, my constant companion who was by my side until the bitter end.  	
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    Chapter 1 Introduction 	
   1.1  Organization  of  the  mammalian  seminiferous  epithelium  and  spermatogenesis Upon examination of a cross section of mammalian testes, two compartments can be observed – the seminiferous tubules and a surrounding interstitium. The interstitium contains blood vessels, lymphatics, connective tissue and Leydig cells (Dym and Fawcett, 1970). The seminiferous tubules contain a unique epithelium that is where the process of spermatogenesis takes place. The seminiferous epithelium is made up of two types of cells – Sertoli cells and developing germ cells. Sertoli cells provide structural and functional support to germ cells throughout development and span the length of the epithelium from basement membrane to tubule lumen (Russell, 1979c). Germ cells lie between Sertoli cells and maintain attachment to them despite their complex arrangement in concentric layers within the tubules (Dym and Fawcett, 1970). The process of spermatogenesis is cyclic and begins as diploid spermatogonia undergo mitosis. After spermatogenesis is initiated, processes of two neighboring Sertoli cells form below spermatocytes and concurrently, move the spermatocyte toward the adluminal compartment (Russell, 1977b). Leptotene spermatocytes translocate through a basal junction complex into the adluminal compartment where they complete meiosis and become haploid spermatids. As development of spermatids progresses, excess cytoplasm is removed in  	
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    preparation for release from the epithelium as spermatozoa (O'Donnell et al., 2011).  1.2 Arrangement and functional significance of intercellular junctions in the seminiferous epithelium The turnover of massive intercellular junctions in the seminiferous epithelium occurs at specific times during spermatogenesis and is associated with two processes that are fundamental to fertility. The first process is the movement of spermatocytes through the basal junction complex and the second process is the release of mature spermatids from the apex into the lumen of the epithelium. At basal sites of attachment between Sertoli cells, the junctions form a complex of tight, gap, desmosome-like and tissue specific adhesion junctions called ectoplasmic specializations. Tight junctions within the basal junction complex form the blood-testis barrier (Fawcett and Dym 1970). At apical sites, the junctions consist almost entirely of ectoplasmic specialization and form between Sertoli cells and late spermatids. The turnover of ectoplasmic specializations represents a major epithelial remodeling event that is specific to the seminiferous epithelium. 1.3 Intercellular junction complexes 	
    	
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    1.3.1 Tight junctions Tight junctions were first described in 1963 as giving epithelial cells the ability to seal off body compartments (Farquhar and Palade, 1963). These junctions exhibit barrier-like properties and are typically found around the circumference of the apical region of epithelial cells (Fig.1.1). The conformation of tight junction components, occludin and claudin, contribute to the paracellular barrier function. Although the two proteins share no sequence similarity, both occludin and claudin contain four transmembrane domains and two extracellular loops (Furuse et al., 1998; Furuse et al., 1993). Stabilization of occludin and claudin is facilitated by scaffold proteins such as ZO-1, which links the junction components to the actin cytoskeleton (Tsukita et al., 2009). The selective permeability of tight junctions plays a functional role in many types of epithelia. A principal tight junction is found at the interface between the brain and blood. In contrast to the blood-brain-barrier, where tight junctions line the blood vessels, the primary component of the blood testis barrier is the tight junctions formed between adjacent Sertoli cells at the basal junction complex (Mital et al., 2011).  1.3.2 Adherens junctions Adherens junctions (AJs) are cell-cell adhesion complexes that are found in most epithelial cells. These dense plaques are intimately associated with the actin cytoskeleton and function to maintain tissue structure and endure various  	
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    stresses. Zonula adherens, a prominent adherens junction, contributes to the formation of a continuous belt that separates epithelial membranes into apical and basolateral compartments (Harris and Tepass, 2010) (Fig. 1.1). Typically, a cadherin – catenin complex make up the core of adherens junctions (Pokutta and Weis, 2007). Cadherins are homophilic, Ca2+ dependent adhesion receptors that consist of two or more extracellular domains and a cytoplasmic tail. Catenins are cytoplasmic adapter proteins that link cadherins to the cytoskeleton. In addition to cadherins, a second class of adhesion receptors, nectins, exists at AJs. Nectins are Ca2+ -independent immunoglobulin-like adhesion molecules that bind to actin via adapter protein afadin (Harris and Tepass, 2010). The cadherin-catenin and nectin-afadin complexes have been shown to behave similarly in mammalian tissue culture studies (Takai et al., 2008; Takai et al., 2003). Each nectin molecule is able to form a homotypic junction with the same molecule of the neighboring cells to mediate cell-cell adhesion. Nectin molecules are also capable of forming heterotypic junctions with other nectin molecules resulting in much stronger interactions (Ozaki-Kuroda et al., 2002).  1.3.3 Gap junctions Gap junctions are specialized intercellular channels that permit direct cellcell transfer of small molecules and ions (Fig. 1.1). Gap junctions are best known for their role in excitable cells, however they are ubiquitous in nearly all solid tissue cells (Goodenough and Paul, 2009). A gap junction channel is composed  	
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    of two connexons that attach across the intercellular space. A connexon is made up of hexamers of integral proteins termed connexions. Like integral tight junction proteins, connexions contain four transmembrane domains with two extracellular loops. A connexon can maintain an open or closed configuration, a characteristic that is vital to a number of cell processes that range from cell coupling to preventing the spread of bacterial infection. Open connexons allow adjacent cells to coordinate responses to various signals, for example, electric conduction within cardiac muscle fibers that form a syncytium (Weidmann, 1952). In the case of bacterial infection, the connexons in enterocytes can loose the ability to maintain a closed configuration resulting in severe dehydration (Guttman et al., 2010). A number of other pathologies are attributed to too few or too many gap junctions in epithelia.  	
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    1.3.4 Desmosomes Desmosomes are a type of intercellular junction typically arranged on the lateral sides of epithelial plasma membranes that tether intermediate filaments to the plasma membrane (Fig.1.1). In contrast to the previously mentioned intercellular junctions, desmosomes have a relatively large intercellular space (30 nm) (Green et al., 1990). Desmosomes anchor intermediate filaments (keratin or vimentin) to adhesion molecules desmogleins, desmocollins, and desmosomal cadherins via adaptor proteins. Desmosomes provide tissues with mechanical strength and integrity and have been implicated in signaling pathways that promote differentiation and morphogenesis (Yang et al., 2006). In addition to maintaining stable cell-cell adhesion, emerging evidence indicates that desmosomes are also dynamic structures that contribute to cellular processes beyond that of cell adhesion (Sumigray et al., 2011).  1.4 Intercellular junctions in the seminiferous epithelium In the basal compartment, germ cells in the early stage of meiosis maintain an attachment to the basal lamina (Russell, 1977b). Sertoli cell processes form below the cells and facilitate a progressive movement of the cells from the basal to the adluminal compartment (Russell, 1977b). Contact between Sertoli cell processes and germ cells as this stage is also facilitated by desomosome-like junctions (Russell, 1977a). Sertoli cells, themselves, are attached to the basal lamina via hemidesmosomes (Russell, 1977b).  	
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    Eventually the Sertoli cell processes that form below spermatocytes meet and form an adherens-like junction. As the spermatocyte moves toward the lumen, contact between the Sertoli processes increases and tight junction contacts are readily visible at this point (Russell, 1977b). This junction complex consists of adherens (ectoplasmic specializations), tight and gap juctions and includes desmosome-like components as well at basal sites (Vogl AW, 2008). At apical sites, ectoplasmic specializations occur between Sertoli cells and germ cells and are made up mostly of adherens junction components.  1.4.1 Ectoplasmic specializations Ectoplasmic Specializations are tripartite structures consisting of Sertoli cell plasma membrane, a layer of hexagonally packed F-actin filament bundles and a cistern of endoplasmic reticulum (Grove and Vogl, 1989). The actin filaments are oriented parallel to the plasma membrane and are linked to both the plasma membrane and the endoplasmic reticulum (Grove and Vogl, 1989). Ectoplasmic specializations have been described as testis-specific, adherensbased junctions (Lee and Cheng, 2004). Ectoplasmic specializations somewhat resemble zonula adherens, a prominent adherens junction that contributes to the formation of a continuous belt that separates epithelial membranes into apical and basolateral compartments (Harris and Tepass, 2010) (Fig. 1.2). Adhesion at ectoplasmic specializations is facilitated by a nectin-afadin complex (Takai and Nakanishi, 2003), although N-  	
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    Cadherin has been reported to be present as well (Yan and Cheng, 2005). The junction molecules in the Sertoli cell plasma membrane at ectoplasmic specializations are nectin 2 (Ozaki-Kuroda et al., 2002) and α6β1 integrin (Salanova et al., 1998). As mentioned previously, each nectin molecule is able to form a homotypic junction with the same molecule of the neighboring cells to mediate cell-cell adhesion. This interaction is observed between two nectin 2 molecules at basal ectoplasmic specializations (Guttman et al., 2004b). The binding partner for nectin 2 in the spermatid plasma membrane at apical sites is nectin 3, forming a hetero-trans-dimer (Satoh-Horikawa et al., 2000). At apical and basal sites, the ligand for α6β1 integrin is reported to be γ3 laminin (Siu and Cheng, 2004), and that for N-cadherin is presumably another Ncadherin molecule in the adjacent membrane, although the presence of Ncadherin at ectoplasmic specializations is controversial. Although ectoplasmic specializations that occur between Sertoli cells and germ cells are made up mostly of adherens junction components, tight junction proteins CAR and JAM-C have been reported to be present (Yan et al., 2007). In addition to these tight junction proteins, intermediate filament – associated protein carcinoembryonic antigen-related cell adhesion molecule 6 (Ceacam6) has been localized to apical ectoplasmic specializations (Kurio et al., 2008).  	
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    1.5 Junction turnover in epithelia Adhesion and tight junction turnover in epithelia generally involves the disengagement of junction molecules of one cell from those on the adjoining cell. Junction molecules are then internalized by a number of mechanisms including clathrin- or caveolin-mediated endocytosis (Akhtar and Hotchin, 2001; Baum and Georgiou, 2011; Ivanov et al., 2004; Izumi et al., 2004; Le et al., 1999; Shen and Turner, 2005; Troyanovsky et al., 2006) and clathrin-/caveolin-/lipid-raft dependent mechanisms such as macropinocytosis (Utech et al., 2005). The multiple pathways of junction endocytosis are dependent upon cell context and regulation (Paterson et al., 2003) but typically involve integral adhesion molecules from one cell detaching from the adhesion molecules in the adjacent cell and being internalized separately. Not all intercellular junction internalization occurs by this method. The internalization of gap junctions occurs via clathrinmediated endocytosis (Piehl et al., 2007) and involves intact gap junctions consisting of connexions in the plasma membranes of the two adjacent cells being internalized as double-membrane vesicles by one of the cells (Piehl et al., 2007).  1.5.1 The endosomal network Once junction material has been internalized, it enters the endosomal network of the cell. In general, the endosomal network is a complex system of vesicles and tubular structures that mediate the trafficking of internalized cargo.  	
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    The first endosome encountered by cargo is the early endosome. The early endosome has a unique geometry with structural elements that are both tubular and vesicular (Marsh et al., 1986). Cargo in the tubular region is recycled back to the plasma membrane. Cargo in the vesicular region will stay in the endosomal system and is likely degraded (Mercer et al., 2010). Early endosomes fuse with late endosomes, also known as multivesicular bodies. There is a significant drop in pH as cargo advances from early to late endosomes. Cargo will undergo an ubiquitin-tagging system in late endosomes called the ESCRT complex. At this stage, cargo is either transported to recycling endosomes for delivery back to the plasma membrane, or to lysosomes for degradation. Internalized intact gap junctions, for example, can be targeted to lysosomes or be recycled (Berthoud et al., 2004; Gaietta et al., 2002). The complex and sophisticated interconnection of the endosomal network has been demonstrated in perturbation assays (Mercer et al., 2010). Rab proteins and their effectors coordinate the communication between endosomal compartments. They are responsible for vesicle formation, motility, and tethering of vesicles to their target compartment. Rabs associate exclusively with specific vesicular membranes. These characteristics make Rabs and other endocytic markers like EEA1 and LAMP1 excellent determinants of endosome identity. Rabs are able to negotiate the challenges associated with endosomal trafficking due to a highly conserved molecular switch mechanism. A  	
    10	
    conformational change takes place when Rabs cycle between GTP-bound (active) and GDP-bound (inactive) conformations (Zerial and McBride, 2001). Different functions are performed depending on the conformational state. A number of effectors facilitate these changes. Anchoring to membranes is facilitated by prenylation (Leung et al., 2006). GDP-bound Rabs transfer between membranes by a Rab GDI (GDP dissociation inhibitor) (Goody et al., 2005). A number of guanine nucleotide exchange factors (GEFs) catalyze exchange from active to inactive conformations (Grosshans et al., 2006). Rabs are deactivated through GTP hydrolysis – a process that is aided by GAPs (GTPase activating proteins) (Bernards, 2003).  1.6 Tubulobulbar complexes Junction internalization in the seminiferous epithelium differs from all of the previous examples. Here, subcellular, actin-based machines termed tubulobulbar complexes internalize ʻintactʼ intercellular junctions into Sertoli cells thereby facilitating the translocation of spermatocytes from basal to adluminal compartments and the release of mature spermatids from the apex of the tubule into the lumen (Russell, 1979a). Russell and Clermont first described tubulobulbar complexes from morphological studies in 1976. The authors report the presence of several long (2-3 m) and narrow (50 nm) tubular projections of the spermatid's plasma membrane that evaginate into the Sertoli cell cytoplasm in the concavity of the  	
    11	
    spermatid head. Tubulobulbar complexes consist of blind-ended tubular projections that extend into Sertoli cells from junctions with adjacent cells. “Bristle-coated” pits occur at the tips of the structures, and distal parts of the complex expand to form bulbar regions (Russell and Clermont, 1976a) (Fig. 1.3, 1.4A). This swollen region lacks an associated network of actin filaments and is closely associated with cisternae of endoplasmic reticulum (Fig. 1.3, 1.4C). The bulbar region eventually buds from the complex and is internalized (Russell and Clermont, 1976a; Russell, 1979b). In the Sertoli cytoplasm surrounding the tubular portion of the structure there is an accumulation of filamentous actin (Russell and Clermont, 1976a) (Fig. 1.3, 1.4B).  1.6.1 Dendritic actin assembly Dendritic actin assembly describes the properties of the leading edge in motile cells. Rather than actin treadmilling, each filament is nucleated at a branch point of Arp 2/3 and a mother filament. A characteristic 700 angle is formed as new actin monomers polymerize at the barbed end. The growth of each filament is transient. Actin capping terminates growth followed by filament severing. Dissociated filaments disassemble and recycle back to the pool of actin monomers to form new filaments (Pollard and Borisy, 2003). This continual polymerization, capping and recycling mechanism generates the force that is necessary to remodel the membrane. Evidence of tubulobulbar complex dependence on dendritic actin assembly was illustrated in an experiment using  	
    12	
    cytochalasin D, an inhibitor of actin polymerization (Russell et al., 1989). In the presence of cytochalasin D, tubulobulbar complexes were unable to develop lacking both the tubular and bulbous portions (Russell et al., 1989).  1.6.2 Basal tubulobulbar complexes Tubulobulbar complexes that form at the basal junction complex consist of a projection of Sertoli cell plasma membrane into a neighboring Sertoli cell. The double membrane tube is cuffed by actin filaments and at the tip is a coated pit (Russell and Clermont, 1976a). In the distal third of basal tubulobulbar complexes, the actin cuff disappears and the tube widens to take on a bulbar configuration that is closely associated with cisternae of endoplasmic reticulum (Russell and Clermont, 1976a). This bulbar region eventually buds off of the complex and enters the Sertoli cell (Russell, 1979a). Basal tubulobulbar complexes form prior to translocation of spermatocytes from the basal to the adluminal compartment (Du et al., 2013; Russell, 1979a) (Fig. 1.2). During the translocation event, the basal junction complex consisting of ectoplasmic specialization, tight and gap junctions disassemble above spermatocytes and simultaneously assemble below (Russell, 1977b). A recent in vitro study of basal tubulobular complexes suggests a role in internalization of intact basal junction components (Du et al., 2013). In addition to basal junction components, nectin-2, claudin-11 and connexin 43 being localized to basal tubulobulbar complexes (Du et al., 2013; Russell, 1979a), endocytic markers Rab5 and EEA1 were found to  	
    13	
    be situated in similar locations (Du et al., 2013), an observation that is consistent with junction internalization. Also, less actin (ectoplasmic specialization) and claudin-11 (tight junction) were present in Stage VII tubules versus Stage V tubules (Du et al., 2013), an observation that correlates with the time that basal tubulobular complexes are known to be most active during spermatogenesis.  1.6.3 Apical tubulobulbar complexes At apical sites, tubulobulbar complexes are made up of a projection of germ cell plasma membrane, originating from regions overlying the acrosome, into the adjacent Sertoli cell (Fig. 1.3). Like basal tubulobulbar complexes, apical tubulobulbar complexes consist of a double membrane tubule with a bulbar region and a coated pit. The tubular region is also cuffed by an actin network and the bulbar region is associated with cisternae of endoplasmic reticulum. Apical tubulobulbar complexes are easier to visualize because of the ability to mechanically fragment the apical Sertoli process. Also, the complexes form in the concavity of the hook shaped head of rat spermatids that provides an easily identifiable fiduciary marker (Fig. 1.1). In comparison to basal tubulobulbar complexes, apical complexes are more numerous, longer, and have a more predictable location (Young et al., 2009b). In contrast, basal complexes are notoriously difficult to visualize due to a lack of fiduciary markers and their tendency to form in junction folds and pockets (Russell, 1979c).  	
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    The focus of this thesis is on the characterization of apical tubulobulbar complexes. The formation of each apical tubulobulbar complex is initiated in the Sertoli cell by a coated pit, which retains an attachment to the neighboring germ cell plasma membrane as the complex elongates and matures (Russell and Clermont, 1976a). In regions of the coated pit where the germ cell plasma membrane remains attached to the Sertoli cell, fine filamentous connections occur between the two cells, and the germ cell membrane contains a distinct submembrane density (Fig. 1.4E). Within an apical Sertoli projection, as many as twenty-four (each up to 2-3 micrometers in length) tubulobulbar complexes develop in two rows.  1.7 Morphologically similar structures Although tubulobulbar complexes are unique to the seminiferous epithelium, they are similar in basic morphology and in molecular composition to a number of other actin-based structures including clathrin-mediated endocytosis machinery (Taylor et al., 2011), to podosomes in osteoclasts (Ochoa et al., 2000) and to membrane-tubules formed in cell free systems (Wu et al., 2010).  	
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    1.7.1 Clathrin-mediated endocytosis machinery Clathrin-mediated endocytosis is a fundamental cellular process of internalization. The basic concepts of clathrin-mediated endocytosis have been understood for decades as researchers have used a combination of electron microscopy and biochemical studies to observe and evaluate the process (Roth and Porter, 1964). Characteristics such as coated pit formation, membrane bending and budding have long been evident but more recent genetic and biochemical studies have been able to shed light on the molecular machinery and mechanisms that are involved. It has become more apparent that clathrinmediated  endocytosis  is  an  incredibly  dynamic  process  involving  the  orchestration of over fifty proteins that interact with each other and the plasma membrane. Live cell imaging of fluorescently tagged proteins has provided insight into the behavior of individual proteins, allowing researchers to organize them into functional modules. The early module, including clathrin and associated coat proteins, initiates assembly (Stimpson et al., 2009). Following the early module, actin recruitment and polymerization increase membrane invagination and finally, scission module proteins facilitate separation of the vesicle from the plasma membrane. Further advances in imaging, genetic and biochemical analysis offer an increasingly detailed understanding of clathrin-mediated endocytosis. For example, live cell TIRF and spinning disc confocal microscopy has been able to capture how a clathrin lattice is initiated. By tagging clathrin triskelion molecules with EGFP, Coccuci et al found that clathrin coat nucleation  	
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    likely begins with one clathrin triskelion binding two AP-2 molecules that were already bound to the plasma membrane (Cocucci et al., 2012). An equally important finding has recently challenged the paradigm that clathrin triskelions and associated coat proteins have the ability to bend the membrane. Kukulski et al. found that membrane curvature does not begin until after actin recruitment and polymerization (Kukulski et al., 2012). Remaining questions about the intricate mechanisms of clathrin-mediated endocytosis will no doubt be answered as these research tools and technologies continue to improve.  1.7.2 Podosomes formed by osteoclasts Podosomes are actin-based structures formed by osteoclasts that develop at sites of cell/substrate attachment and consist of elongate tubular plasma membrane cores surrounded by cuffs of actin filaments (Ochoa et al., 2000). Podosomes and tubulobulbar complexes have a number of molecular components in common and both structures involve the dendritic assembly of actin in their assembly. Although podosomes are not clathrin-mediated, their formation is generated by actin network proteins N-WASP, Arp2/3, cortactin, and dynamin in addition to BAR proteins (Bharti et al., 2007; Buccione et al., 2004; Ochoa et al., 2000; Tehrani et al., 2006). BAR (Bin-Amphiphysin-Rvs) proteins are highly conserved and are involved in membrane bending (Wu et al., 2010). The function of podosomes is not entirely clear although adhesion and matrix degradation are likely (Buccione et al., 2004).  	
    17	
    1.7.3 Reconstituted membrane tubules Clathrin-mediated bulk endocytosis is a description that was coined in reference to the behavior of reconstituted membranes in a cell free system (Wu et al., 2010). When fibroblast plasma membranes are incubated in cytosol with ATP and a G-protein analog, clathrin-mediated tubules form. These 50-120 nm tubules are formed through the coordinated action of a BAR domain protein, clathrin and actin (Wu et al., 2010). Dynamin labeled a narrowed, distal portion of the tubules (35nm) near the clathrin-coated pit. The BAR protein labeled the proximal region of the tubule that was characteristically wider (120nm) than the distal region. The generated tubules were actin dependent and failed to form upon addition of latrunculin to the system. When latrunculin was added to the system after tubule formation, the actin cytoskeleton was depleted, but the tubules remained intact. These results suggest that as in tubulobulbar complex formation, actin aids in tubule formation of the membrane tubule, while the FBAR protein acts as a scaffold to support tubules (Wu et al., 2010). Podosomes formed by osteoclasts, reconstituted membrane tubules, the necks of mature clathrin coated-pits and tubulobulbar complexes share morphological similarities in that they all consist of elongate cores of tubular membrane surrounded by networks of actin filaments and related proteins. However, the core of tubulobulbar complexes consists of the plasma membranes  	
    18	
    of two adjacent cells whereas the other structuresʼ central core is a single membrane.  1.8 Proposed functions of tubulobulbar complexes The location, timing and unique structure of tubulobulbar complexes have lead to a number of suggested functions. One proposal is that tubulobulbar complexes eliminate cytoplasm from spermatids and reduce their volume prior to release (Russell, 1979d). This a sensible proposal based on the observation that in rat, there is a 50% reduction in the volume of cytoplasm during the formation of tubulobulbar complexes (Sprando and Russell, 1987). Another hypothesis is that tubulobulbar complexes are simply attachment devices that firmly link elongate spermatids to the seminiferous epithelium during the dynamic process of spermatogenesis (Russell, 1979b). One groupʼs suggestion is that tubulobulbar complexes facilitate the shaping of “hooked” spermatid heads in rat (Kierszenbaum and Tres, 2004), and remove excess acrosomal contents from spermatids (Tanii et al., 1999).  	
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    1.9 Working hypothesis and chapter aims The incompatibility with all of these functions is that they focus on Sertoli cell interactions with spermatids and do not account for the formation of tubulobulbar complexes at the basal junction complex. The working hypothesis of this thesis is that tubulobulbar complexes are subcellular machines that internalize intercellular junctions and are at least partially responsible for spermatid release at apical sites in the seminiferous epithelium and spermatocyte translocation at basal sites (Du et al., 2013; Russell, 1979c; Russell et al., 1988; Vogl, 1989). This hypothesis is consistent with the use of clathrin-based endocytic machinery and the timing of tubulobulbar complex formation prior to junction remodeling events in the seminiferous epithelium. In animals that have been treated with estradiol (D'Souza et al., 2009) and in amphiphysin knock out animals, tubulobulbar complexes fail to form (Kusumi et al., 2007). This phenotype is correlated with failure of spermatids to be released from the epithelium (D'Souza et al., 2009; Kusumi et al., 2007). Also, spermiation failure is one of the first phenotypes observed in men undergoing hormonal methods of contraception (McLachlan et al., 2002). Members of the Vogl lab have previously shown evidence of junction internalization (Guttman et al., 2004b). Nectin 3, the binding partner of nectin 2 that is expressed by germ cells at ectoplasmic specializations, is absent from spermatozoa that have been released from the epithelium (Guttman et al., 2004b). Significantly, both nectin 3 and nectin 2 have been localized to the distal  	
    20	
    ends of tubulobulbar complexes – an indication of internalization (Guttman et al., 2004b). Furthermore, antibodies to endosomal marker LAMP1 and lysosomal marker SGP1 label the cluster of vesicles associated with the ends of tubulobulbar complexes (Guttman et al., 2004b). In this thesis, I explore the prediction that tubulobulbar complexes are responsible for the internalization of intercellular junctions. Four main predictions about tubulobulbar complexes are addressed in the following chapters. The first prediction is that if structures that morphologically resemble tubulobulbar complexes are formed by a similar cytoskeletal network, then key components found at these sites also should be present at tubulobulbar complexes. Previous members of the Vogl lab have identified dendritic actin assembly components Arp2/3 and dynamin 3 (Vaid et al., 2007) to be present at tubulobulbar complexes. In the first chapter of this thesis, I test the prediction that N-WASP and cortactin also are present at these sites. N-WASP and cortactin are two key components of dendritic actin assembly that likely contribute to tubulobulbar complex formation. Arp 2/3 is inactive on its own and completely dependent on nucleation promoting factors like the WASP/Scar family of proteins to bind to actin (Higgs and Pollard, 2000). Until activated, N-WASP exists in its auto-inhibited state due to its GTPase domain folding upon itself and binding its cofilin-homology sequence. Upon binding of GTPase Cdc42 and PI(4,5)P2, NWASP unfolds into its active form. Once in itʼs open, activated state, N-Waspʼs Cterminal VCA domain that is responsible for Arp 2/3 binding and actin nucleation  	
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    becomes accessible and dendritic actin assembly begins (Machesky and Insall, 1998; Marchand et al., 2001; Miki and Takenawa, 1998). Cortactin plays a dynamic role in actin binding. Cortactin monomers, diffuse through the cytoplasm, contain actin binding and SH3 domains that contribute to the proteinʼs dynamic capabilities. Extracellular signals cause phosphorylation of cortactin that subsequently binds and recruits Arp2/3 to existing actin filaments. Cortactin functions to stabilize newly forming actin branches and inhibits disassembly (Daly, 2004). The localization of N-WASP and cortactin to tubulobulbar complexes will contribute to the understanding of their formation. If tubulobulbar complexes are generated by the assembly of an actin network involving the action of N-WASP and cortactin in addition to Arp2/3 and dynamin 3, then it will be possible to include dendritic actin assembly in the model of tubulobulbar complex formation. Also in this chapter, I explore the possibility that clathrin is localized to the coated pits at the ends of tubulobulbar complexes. The presence of clathrin at tubulobulbar  complexes  would  suggest  formation  by  clathrin-mediated  endocytosis. Chapter three focuses on the localization of the structural components zyxin and vinculin. Both proteins are elements of focal adhesions and during simultaneous events of junction disassembly and podosome formation, zyxin and vinculin redistribute to podosomes and co-localize at these sites (Kaverina et al., 2003). Based on the behavior of zyxin and vinculin at focal adhesions and  	
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    podosomes in smooth muscle, I test the prediction that zyxin and vinculin – previously localized to ectoplasmic specializations – co-localize at tubulobulbar complexes. In chapter four, I address the second of the four main predictions by expanding on the previous findings that adhesion proteins nectin 2 and nectin 3 are localized to tubulobulbar complexes. If tubulobulbar complexes do internalize intact intercellular junctions, then junction material should not only be present at tubulobulbar complexes but should also be associated with endosome markers and the ends of the complexes. Also, α6 integrin, an integral adhesion molecule known to be present at ESs, should be present at tubulobulbar complexes. To test this hypothesis, I probe testis sections and fragments for early endosome antigen 1 (EEA1) and attempt to confirm previous reports that N-cadherin is present at ectoplasmic specializations. In the fifth chapter, I test the third prediction about tubulobulbar complexes. If tubulobulbar complexes internalize regions of intact intercellular junction, then junction material must enter the endosomal network of Sertoli cells. In this chapter, I explore the possibility that the β1 integrin subunit that is reported to be present at ectoplasmic specializations is present at tubulobulbar complexes (Salanova et al., 1995). The presence of β1 integrin at tubulobulbar complexes would be consistent with the junction internalization hypothesis. I also probe rat seminiferous epithelium fragments and sections for nectin 2 and nectin 3 – adhesion proteins that have previously been localized to tubulobulbar complexes  	
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    in mouse. This chapter also turns the focus to the vesicles at the ends of tubulobulbar complexes and aims to gain a deeper understanding of the fate of junction material after it enters the endosomal network. The trafficking of vesicles associated with tubulobulbar complexes in Sertoli cells remains to be analyzed. The contents of these vesicles could be recycled back to the plasma membrane or targeted for degradation in lysosomes.  This chapter reports the findings  of the extensive probing of the Sertoli cellʼs endosomal network with various endocytic markers to aid in the understanding of junction molecule trafficking and fate. Chapters two - five of this thesis have included a three-fold approach to characterizing tubulobulbar complexes. The objective has been to gain insight into how tubulobular complexes are formed, if integral junction material is present within them, and the nature of the associated vesicles. These descriptive studies have gathered vital information about tubulobulbar complexes making it possible to build a model of tubulobulbar complex formation, junction internalization and junction degradation/recycling. Chapter six of this thesis tests the last of the four predictions about tubulobulbar complexes. In this chapter, the models that were generated by the work in previous chapters are utilized to employ functional experiments that test the prediction that if the structure/function of tubulobulbar complexes is perturbed, then sperm release will be delayed. In these experiments, an intratesticular approach was used to knockdown the expression of key actin-  	
    24	
    related protein cortactin. Using this method, cortactin-targeting siRNA was injected into testis to determine if the knockdown had an effect on tubulobulbar complexes. If tubulobulbar complexes are responsible for internalizing intact intercellular junctions, then reducing the amounts of the actin-binding component cortactin should interfere with their structure/function as occurs in podosomes in cultured osteoclasts (Tehrani et al., 2006). Perturbation of tubulobulbar complex function should result in the delay of sperm release. Observation of the expected phenotype from this work will ultimately characterize the biological function of tubulobulbar complexes.  	
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    1.10 Introduction figures  Figure 1.1 Intercellular junctions in epithelia in general. Four types of cell-cell junctions are represented in this schematic diagram of intestinal epithelial cells. Tight junctions (red) are impermeable and prevent molecules from passing through the intercellular space. Adherens junctions (green) form belt-like adhesions utilizing actin to bind adjacent cells. Gap junctions (purple) have the ability to connect the cytoplasm of two adjacent cells. Desmosomes (orange) facilitate cell adhesion through the use of intermediate filaments. Modified from(Itoh and De Camilli, 2006)Nature Reviews Molecular Cell Biology.  	
    26	
    Figure 1.2 Schematic diagram showing a section of seminiferous epithelium. Sertoli cells are in light gray and spermatogenic cells are in a darker gray. The head of a late spermatid (black) is illustrated at the apex of the epithelium. Ectoplasmic specializations (ES) occur in apical regions between Sertoli cells and spermatids and at basal junction complexes between neighboring Sertoli cells.  	
    27	
    Figure 1.3 Diagram of the structure of tubulobulbar complexes in the seminiferous epithelium. Basal tubulobulbar complexes occur in association with junctions between two neighboring Sertoli cells. Apical complexes occur in regions where Sertoli cells are adherent to spermatids. Each tubulobulbar complex consists of an elongated invagination into a Sertoli cell of the plasma membranes of two cells at the site of intercellular junction. The complex consists of tubular and bulbar regions. A coated pit occurs at the end of the complex. The proximal tube has a diameter of 50 nm and has a double membrane. Vesicles budding from the distal tube also consist of a double membrane. Modified from (Vaid et al., 2007) Journal of Cellular Physiology.  	
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    Figure 1.4 Ultrastructural features of tubulobulbar complexes associated with Sertoli/spermatid attachment sites at the apex of rat seminiferous epithelium. (A) Longitudinal section of a tubulobulbar complex. Bar = 200 nm. (B) Cross section through proximal tubular region. Bar = 100 nm. (C) Cross section through bulbar region. Bar = 100 nm. (D) Longitudinal section through distal tubular region and coated-pit. Notice the density (asterisk) associated with the spermatid plasma membrane and the connections (arrowhead) with the Sertoli cell plasma membrane of the coated pit. Bar = 100 nm. (E) Cross section through a coated pit illustrating fine filamentous attachments between the plasma membranes of the Sertoli cell and the spermatid (arrowheads), and the density associated with the spermatid plasma membrane (asterisk). Bar = 100 nm.  	
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    Chapter 2 – Cortactin (CTTN), n-wasp (WASL) and clathrin (CLTC) are present at podosome-like tubulobulbar complexes in rat testis  2.1 Brief synopsis Although the basic structure of tubulobulbar complexes is well established, few of their molecular components have been identified previous to the publication of this chapter. Tubulobulbar complexes resemble podosomes in other systems (Goicoechea et al., 2006; Ochoa et al., 2000). Podosomes that occur in osteoclasts form at sites of cell/substrate attachment and consist of elongate tubular plasma membrane cores surrounded by cuffs of actin filaments (Ochoa et al., 2000). Podosome formation is generated by the assembly of an actin network involving the action of N-WASP (WASL), the Arp2/3 complex, cortactin (CTTN), and dynamin (DNM) (Buccione et al., 2004; Ochoa et al., 2000; Tehrani et al., 2006). Previous members of the Vogl lab had localized Arp2/3 and dynamin 3 to tubulobulbar complexes (Vaid et al., 2007), and others have reported dynamin 2 and amphiphysin (Kusumi et al., 2007), a membrane-curvature sensing protein, to be present at the structures. In this study, the prediction that two key molecular components, N-WASP and cortactin, found at podosomes also are present at tubulobulbar complexes was explored as well as the possibility that the bristle-coated pits at the ends of  	
    30	
    tubulobulbar complexes contain clathrin. Rat testis was used as a model system because tubulobulbar complexes are very prominent and are best studied in this species. Apical tubulobulbar complexes that form in association with late spermatids were the focus because these complexes are more numerous, longer, and more predictable in location than are basal tubulobulbar complexes. Also, late spermatids together with adjacent Sertoli cell regions can be mechanically dissociated from the seminiferous epithelium and used to clearly resolve  the  tubulobulbar  complexes.  Testis  sections  and  mechanically  dissociated fragments of seminiferous epithelium were probed for cortactin, NWASP, and clathrin. This study has two significant findings. First, cortactin and N-WASP, key components present at podosomes, also are present at tubulobulbar complexes. Second, clathrin is a component of the bristle-coated pit at the ends of the structures. The results from this study are consistent with the general hypothesis that tubulobulbar complexes are podosome-like structures associated with junction turnover in the seminiferous epithelium, and raise the interesting possibility that formation of the structures may be clathrin mediated. I propose a mechanism for the formation of tubulobulbar complexes that incorporates both the dendritic model of actin assembly and clathrin-mediated endocytosis (Kaksonen et al., 2006; Mullins et al., 1998).  	
    31	
    2.2 Results  2.2.1 Tubulobulbar complexes contain n-wasp and cortactin Because tubulobulbar complexes structurally resemble podosomes in other systems, I wanted to determine whether or not key molecular components found at podosomes also are present at tubulobulbar complexes, particularly because many of these components are involved in podosome formation and may provide insight about tubulobulbar complex formation. Among these components are the actin-related protein cortactin and the Arp2/3 activator NWASP.  2.2.2 Tubulobulbar complexes contain cortactin The antibody raised against cortactin and used in these studies reacted on Western blots of seminiferous epithelium and testis in a fashion characteristic for cortactin (Fig. 2.1a) that is, the antibody reacted with multiple bands between 75 and 85 kDa. On frozen sections of fixed testis and on fixed fragments of seminiferous epithelium, diffuse linear tracts of staining were present at basal junction complexes and at sites associated with spermatid heads (Fig. 2.2). Staining at basal junction complexes and at sites associated with spermatid heads, interpreted as being ectoplasmic specializations, has been previously reported and serves as a positive control for the intense staining that was observed in association with tubulobulbar complexes at stage VII of  	
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    spermatogenesis (Fig. 2.1). This was particularly evident in epithelial fragments viewed at high magnification, where cortactin clearly outlined individual complexes adjacent to the concave surface of late spermatid heads (Fig. 2.1f) Controls for staining were negative (Figs. 2.1 and 2.2).  2.2.3 Tubulobulbar complexes contain n-wasp The antibody to N-WASP reacted specifically with a band at the appropriate molecular weight for the protein (65 kDa) on Western blots of testis and seminiferous epithelium that was not present on companion blots labeled with normal rabbit immunoglobulin G (Fig. 2.3) Other reactive bands on the blots were nonspecific and were present on control (normal rabbit IgG) blots. In sections, the N-WASP antibody reacted with ring-like structures related to the cytoplasm of elongate spermatids (Fig. 2.4a) and with residual lobes of late spermatids. Unlike the probe for cortactin, the N-WASP antibody did not react in regions adjacent to spermatid heads nor with junction complexes near the base of the epithelium. However, short elongate profiles were seen occasionally near the base of the epithelium that were likely basal tubulobulbar complexes (Fig. 2.4c) The antibody strongly labeled regions containing tubulobulbar complexes adjacent to the concave surface of late spermatid heads (Fig. 2.4c). In epithelial fragments containing late spermatids, the N-WASP antibody clearly labeled individual tubulobulbar complexes (Fig. 2.5). Patterns of staining seen in  	
    33	
    antibody-treated sections and fragments were not seen in any of the control slides (Figs. 2.4 and 2.5).  2.2.4 Clathrin is localized to the ends of tubulobulbar complexes Bristle-coated pits have been described at the ends of tubulobulbar complexes, and clathrin, a component of coated pits, has been implicated in recruiting the actin cytoskeleton to sites of endocytosis in other systems (Kaksonen et al., 2005). To determine whether clathrin is present at tubulobulbar complexes, rat testis samples were probed with a clathrin antibody using immunofluorescence and immunogold techniques. The antibody reacted with a band of the appropriate molecular weight (180 kDa) for clathrin heavy chain on Western blots of rat testis and seminiferous epithelium (Fig. 2.6a) and also reacted with small punctate structures in epithelial fragments processed for immunofluorescence and that contained apical tubulobulbar complexes associated with late spermatids (Fig. 2.6 b, and d). In some fragments, these punctate structures were localized near where the ends of tubulobulbar complexes are known to occur (Fig. 2.6b), whereas in other samples the punctate profiles were more scattered in their arrangement (Fig. 2.6d). Similar patterns were not seen in control samples treated with normal mouse IgG instead of the primary antibody, even though there was significant diffuse background staining present in these samples (Fig. 2.6, c and d). No  	
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    staining was present in controls for the secondary antibody (Fig. 2.6f) or for autofluorescence (Fig. 2.6g). In thin sections of rat testis processed for clathrin localization by immunogold electron microscopy, discrete clusters of gold particles were seen associated with Sertoli cell regions containing apical tubulobulbar complexes (Fig. 2.6 h and i). In favorable sections, some of these clusters situated at the ends of the complexes (Fig. 2.6 h and i). A similar staining pattern was not present in material treated with normal mouse IgG (Fig. 2.6j). In these control sections, there was a diffuse distribution of gold particles in regions associated with tubulobulbar complexes. In secondary antibody control (2.6k), gold particles were virtually absent from similar regions.  2.3 Discussion In this study, N-WASP and cortactin were demonstrated to be concentrated at intercellular junction-related tubulobulbar complexes in the seminiferous epithelium of the testis. Based on these results, it is logical to propose that the formation of tubulobulbar complexes likely involves the dendritic assembly of an actin network and may be clathrin mediated. Tubulobulbar complexes at intercellular junctions in the seminiferous epithelium structurally resemble podosomes formed at cell substrate adhesion sites in other systems in vitro (Ochoa et al., 2000). Both structures develop at sites of attachment, and both are characterized by a cylindrical column of actin  	
    35	
    filaments that are arranged as a three-dimensional network. Significantly, podosomes in osteoclasts have a tubular core of membrane that invaginates from the plasma membrane, similar to the double plasma membrane core of tubulobulbar complexes in the testis (Ochoa et al., 2000). The function of podosomes is not entirely clear, although roles in adhesion and in matrix degradation are proposed (Buccione et al., 2004). The formation of podosomes is dependent on the cooperative interaction between N-WASP, Arp2/3, cortactin, and dynamin (Buccione et al., 2004; Tehrani et al., 2006). N-WASP is a key activator of the Arp2/3 complex, which in turn generates new actin filament branches from preexisting filaments, thereby forming a three-dimensional network in which filaments elongate from their barbed ends positioned at the plasma membrane. This dendritic actin network assembly is thought to be responsible for the protrusion of lamellipodia at the leading edge of motile cells (Mullins et al., 1998). The dynamins as a group are large GTPases involved with the formation of vesicles and membrane tubules in certain conditions, and with modulating actin dynamics (Gray et al., 2005; McNiven, 1998; Ochoa et al., 2000; Takei et al., 1995). Cortactin is a multidomain, actin-associated protein that binds to actin, the Arp2/3 complex, dynamin, and N-WASP, among others, and is a central regulator of actin filament nucleation and dendritic actin network assembly (Buccione et al., 2004; Weed and Parsons, 2001). Previous members of the Vogl lab have demonstrated that Arp2/3 and Dynamin 3 are present at tubulobulbar complexes  	
    36	
    and this study shows for the first time that N-WASP and cortactin are present as well (Vaid et al., 2007). Results from this and other studies demonstrate that actin filaments, Arp2/3, N-WASP (this study), cortactin (this study), cofilin, espin, dynamin 3, and amphiphysin are present at tubulobulbar complexes, and that clathrin is a component of the coated pits associated with their ends (Guttman et al., 2004a; Guttman et al., 2004b; Kusumi et al., 2007; Vaid et al., 2007; Vogl, 1989). A summary of the localization of some of these components to tubulobulbar complexes is shown in Figure 2.7a, a diagram showing the sequence of events of tubulobulbar complex formation is shown in Figure 2.7b, and micrographs of coated pits at the ends of tubulobulbar complexes are shown in Figure 2.7c. The presence of actin-related components, clathrin, and dynamin in tubulobulbar complexes leads to the proposal of the following model (Fig. 2.8) for tubulobulbar complex formation, which is based on current models of clathrinmediated endocytosis, the dendritic model of actin assembly, and results from studies of podosome formation (Buccione et al., 2004; Kaksonen et al., 2006; Mullins et al., 1998; Pollard, 2007). A clathrin-coated pit formed at junction sites initiates the formation of tubulobulbar complex by recruiting and coupling the actin cytoskeleton to the sites. Unlike in conventional models of endocytosis the clathrin-coated pit remains associated with the forming structure and does not  	
    37	
    detach. Consistent with this possibility is the observation that a coated pit is consistently observed at the end of each tubulobulbar complex and that the pit encloses, and is morphologically attached by fine filamentous connections to, a dense nodular protrusion of the adjacent cell. This clathrin-coated pit may stabilize the tubulobulbar complex, or perhaps even continue to recruit actin machinery and membrane-related components to the site and facilitate tubulobulbar elongation. N-WASP and cortactin activate Arp2/3-mediated dendritic actin network assembly, in which new filaments are established as branches from preexisting filaments, and polymerization of filaments occurs from free barbed ends of the filaments located near the plasma membrane. Dendritic actin assembly is likely the force that drives elongation of the structure. A membrane-associated BAR domain protein (amphiphysin), which senses curvature and bends membranes, and dynamin 3 facilitate formation of the tubular core (Peter et al., 2004). Cortactin may stabilize actin filament branch points and link the network to dynamin at the plasma membrane. Filament crosslinkers, such as α-actinin and espin, may cross-link actin filaments within the forming network. Network turnover is mediated by cofilin, which disassembles the actin filaments, and filament length is controlled by the predicted presence of a barbed-end capping protein. The actin monomer pool available for polymerizing is maintained by profilin, which we predict to be present at the sites. Loss of actin filaments in distal regions of the structures lead to a swelling in the bulbar region, and this structure is associated with, and perhaps its formation is regulated by, a  	
    38	
    close relationship with a cisternae of endoplasmic reticulum. Budding of the bulbar region and eventual vesiculation of the tubulobulbar complex may be generated by the regulated “pinchase” action of dynamin 3. Results of this study support the general conclusion that dendritic actin assembly is involved with tubulobulbar complex formation in the testis, and that the process may be clathrin-mediated.  2.3 Chapter 1 figures  Figure 2.1 Cortactin immunofluorescence and western blot Paired phase and immunofluorescence micrographs of sections and fragments of rat seminiferous epithelium at stage VII of spermatogenesis labeled for cortactin. a) Western blot of testis and seminiferous epithelium labeled with the cortactin antibody. Molecular weights are in kDa. b) Section clearly illustrating an intense signal (arrowheads) associated with the concave face of spermatid heads near the apex of the epithelium. c) Similar section treated with normal rabbit IgG (NRIgG) instead of primary antibody. d) Secondary antibody control. e) Control for autofluorescence. f) Epithelial fragment in which tubulobulbar complexes are clearly labeled with the cortactin antibody (arrow). g) Specificity control treated 	
    39	
    with normal NRIgG instead of primary antibody. h) Control for secondary antibody (2° Control). i) Control for autofluorescence (Blank). Bars = 10 μm.  Figure 2.2 Cortactin immunofluorescence stage V Paired phase and immunofluorescence micrographs of sections and fragments of rat seminiferous epithelium at approximately stage V of spermatogenesis labeled for cortactin. Although ectoplasmic specializations occur in association with elongate spermatids at this stage of spermatogenesis, tubulobulbar complexes have not yet formed. a) The antibody labels regions associated with the heads of spermatids (small arrowheads) and with basal junction complexes (large arrowhead). b) A similar staining pattern is not present in sections treated with normal rabbit IgG (NRIgG) instead of primary antibody. c) Control for secondary antibody. d) Control for autofluorescence. e) Epithelial fragment containing an  	
    40	
    elongate spermatid and associated Sertoli cell ectoplasm specialization. Labeling for cortactin occurs in the ectoplasmic specialization that appears as a linear signal (arrowheads) that outlines the spermatid head. f) Epithelial fragment stained with normal rabbit NRIgG instead of primary antibody. g) Secondary antibody control (2° Control). h) Control for autofluorescence (Blank). Bars = 10 μm (a–d) and 5 μm (e–h).  Figure 2.3 Western blot of rat testis and seminiferous epithelium labeled with a n-wasp antibody and normal rabbit IgG (NRIgG). The antibody specifically labels a band at approximately 65 kDa (asterisk-labeled arrow) that corresponds to the molecular weight of N-WASP. Other reactive bands (kDa) are nonspecific and also appear in the blot labeled with NRIgG.  	
    41	
    Figure 2.4 Paired phase and immunofluorescence micrographs of rat seminiferous epithelium labeled for n-wasp. a) Section of seminiferous epithelium at stage V of spermatogenesis labeled for N-WASP. Staining occurs as ring like or circular profiles (arrows) associated with the cytoplasm of elongate spermatids. Labeling is not concentrated in regions associated with spermatid heads. Although ectoplasmic specializations occur in association with elongate spermatids at this stage of spermatogenesis, tubulobulbar complexes have not yet formed. b) Similar section as in a, but treated with normal rabbit IgG (NRIgG) instead of primary antibody. c) Section of epithelium at stage VII of spermatogenesis. Labeling with the N-WASP antibody occurs in association with tubulobulbar complexes associated with late spermatids (large arrow), short linear profiles at the base of the epithelium (arrowheads), and developing residual lobules (small arrow). d) Control for specificity treated with normal rabbit IgG rather than with primary antibody. Bar = 10 μm.  	
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    Figure 2.5 Paired phase and immunofluorescence micrographs of epithelial fragments of rat seminiferous epithelium containing late spermatids and associated sertoli cell regions labeled for n-wasp. a) The N-WASP antibody labels tubulobulbar complexes (arrowhead). b) Specificity control for the primary antibody. c) Control for secondary antibody (2° Control). d) Control for autofluorescence (Blank). Bars = 5 μm.  	
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    Figure 2.6 Immunofluorescent and immunoelectron microscopic demonstration of clathrin at apical tubulobulbar complexes in the rat. a) Western blots of testis and seminiferous epithelium labeled with a clathrin antibody (left) and normal mouse IgG (NMIgG; right). The clathrin antibody reacts with a specific band at approximately 180 kDa (arrowhead) and a nonspecific band (small asterisk) that also occurs in the blots treated with NMIgG. b–g) Paired phase and fluorescence micrographs of rat seminiferous epithelial fragments labeled for clathrin and control reagents. The clathrin antibody labels  	
    44	
    small punctate structures in regions near where the ends of tubulobulbar complexes are known to occur (arrows in b and d) as well as more generally in the region (d). Similar punctate patterns are not present when the primary antibody is replaced with NMIgG (c and e), when the primary antibody is replaced with buffer alone (2° Control; f), or when both the primary and the secondary antibodies are replaced with buffer alone (Blank; g). h–k) Localization of clathrin at the ultrastructural level. In thin sections labeled for clathrin, gold particles occur in clusters (arrowheads in h and i), some of which appear at the ends of tubulobulbar complexes. When the primary antibody is replaced with NMIgG, gold particles are diffusely present on the section (j). When the primary antibody is replaced with buffer alone (2° Control), few gold particles are present on the section (k). Large asterisks show spermatid heads. Bars = 5 μm (b–g) and 200 nm (h–k).  	
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    46	
    Figure 2.7 Known components and structural features of tubulobulbar complexes together with a diagram showing the formation of the complexes. a) Paired phase (or differential interference contrast) and immunofluorescence micrographs summarizing some of the known components of tubulobulbar complexes. b) Schematic diagram of an apical Sertoli cell process (light gray) with an attached late spermatid (dark gray and black). A number of tubulobulbar complexes are illustrated in a developmental sequence from left to right associated with the concave aspect of the spermatid head. The asterisk indicates the approximate phases of tubulobulbar formation illustrated in Figure 8. c) Electron micrographs showing coated pits at the tips of tubulobulbar complexes just distal to the swollen bulbar regions. In the micrograph on the right, notice that the pit appears attached by fine filamentous connections to the adjacent process of the spermatid, and that there is an electron density at the end of this process. Bars = 5 μm (a) and 200 nm (c).  	
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    Figure 2.8 Model of how dendritic actin dynamics, clathrin coat proteins, and peripheral plasma membrane proteins may interact to generate tubulobulbar complexes. The approximate stage in formation of the tubulobulbar complex illustrated here is indicated by the asterisk in Figure 1.7b. The plus signs (+) indicate the barbed ends of the actin filaments that are directed toward the plasma membrane. Not shown here are the junction molecules that would be in the plasma membranes of the two attached cells.  	
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    Chapter 3 – Focal adhesion proteins zyxin and vinculin are co-distributed at tubulobulbar complexes in rat testis  3.1 Brief synopsis In the previous chapter, I reported that tubulobulbar complexes contain similar actin related components including N-WASP, Arp2/3, cortactin and dynamin 3 to those reported to be present at podosomes. In this chapter, I speculate that the two structures also may have other molecular components in common. One of these components is zyxin. Zyxin is an 82 kDa protein that is known to concentrate at focal adhesions (Yoshigi et al., 2005). It has been described as a mechanosensitive protein due to its ability to mobilize and relocate from focal adhesions to actin stress fibers in response to mechanical cues (Yoshigi et al., 2005). Zyxin is reported to be present at smooth muscle podosomes (Kaverina et al., 2003). In smooth muscle cells, phorbol ester triggers the conversion of focal adhesions into podosomes (Kaverina et al., 2003). At discrete microdomains on the ventral surface of the plasma membrane, podosomes form near the sites where stress fibers insert into adhesion plaques (Webb et al., 2005). These structures are reported to contain α-actinin, F-actin and vinculin and exhibit a tubular, column-like structure arising perpendicularly to the “bottom” of phorbol dibutyrate treated cells (Hai et al., 2002). The composition of smooth muscle  	
    49	
    podosomes has not been confirmed at the ultrastructural level. Despite this lack of morphological evidence, their structure has been included in the description of podosomes that have a central membrane invagination surrounded by an actin filament cuff that in turn is encircled by a larger ring-shaped structure containing focal adhesion proteins such as α-actinin and vinculin (Webb et al., 2005). Also, podosome formation is dependent on Arp2/3-dependent actin polymerization (Kaverina et al., 2003). Smooth muscle cell focal adhesions contain structural proteins zyxin and vinculin (Kaverina et al., 2003). During the simultaneous events of junction disassembly and podosome formation, cytoskeletal rearrangement takes place. At later stages of podosome formation, zyxin and vinculin redistribute to podosomes and co-localize at these sites (Kaverina et al., 2003). Both zyxin and vinculin have previously been reported to be present at ectoplasmic specializations in the seminiferous epithelium (Grove et al., 1990; Lee et al., 2004). Furthermore, vinculin has been reported to be present at tubulobulbar complexes (Kusumi et al., 2007). In this study, we explored the prediction that these two major structural components, vinculin and zyxin, are co-distributed at tubulobulbar complexes. We immunologically probe cryosections and epithelial fragments of perfusion fixed testis for vinculin and zyxin. As predicted, zyxin and vinculin co-distribute at apical tubulobulbar complexes. Rather than the staining of individual complexes that was observed  	
    50	
    with actin assembly proteins, vinculin and zyxin display a more diffuse, although specific association with tubulobulbar complexes.  3.2 Results 	
   3.2.1  The  zyxin  antibody  labels  ectoplasmic  specializations  and  tubulobulbar complexes. In sectioned tissue, the zyxin antibody reacted at tubulobulbar complexes, in addition to reacting with ectoplasmic specializations as previously reported (Lee et al., 2004). Interestingly, the staining varies according to the stage of the tubule that is observed. At stage V of spermatogenesis when spermatids are situated deep in apical Sertoli cell crypts, the zyxin antibody clearly labels ectoplasmic specializations associated with the crypts as indicated by intense staining with phalloidin (Fig. 3.1A). In stage VII tubules, staining is present adjacent to the convex or dorsal face of late spermatids where ectoplasmic specializations are known to occur, and around tubulobulbar complexes clustered adjacent to the convex face of the hook-shaped spermatid heads (Fig. 3.1B). Fragments of seminiferous epithelium viewed at 100X reveal intense staining associated with the dorsal curvature of spermatid heads and diffuse staining around tubulobulbar complexes (Fig. 3.2) although staining of individual complexes also was observed. Phalloidin can be seen labeling the actin cuff around individual complexes (Fig. 3.2).  	
    51	
    3.2.2 Zyxin is present with vinculin at tubulobulbar complexes Zyxin and vinculin both are present at tubulobulbar complexes. The staining pattern for both proteins appears diffuse, and in some cases outlines individual complexes that are adjacent to the concave surface of mature spermatid heads. A Western blot of whole testis and seminiferous epithelium lysate confirms the presence of vinculin (Fig. 3.3). Tubulobulbar complexes are best resolved in fragmented tissue (Young et al., 2009a). Zyxin can be seen diffusely labeling the Sertoli cell cuff in regions where tubulobulbar complexes are present. These staining patterns are absent in controls (Fig. 3.4).  3.2.3 In testis, the zyxin antibody is specific for zyxin splice variant HED-2. The zyxin antibody B71 (Beckerle lab) that was used in these studies reacted on Western blots of whole testis and seminiferous epithelium lysate at the appropriate molecular weight of 82 kDa (Fig. 3.4E). Interestingly, a second, more intense band at 20 kDa was present (Fig. 3.4E). A band of this weight has been previously reported to be present in human seminiferous epithelium and represents zyxin splice variant HED-2 (Yang et al., 1998). Similar gene splicing may be occurring in rat although this remains to be determined. Like full-length zyxin, HED-2 contains a proline rich domain and three LIM domains. HED-2 is present in Sertoli cells and other tissues (Yang et al., 1998). A band also is present at 66 kDa. Hoffman and coworkers have reported this band previously in  	
    52	
    platelets as a protein that may have similarity to zyxin (Hoffman et al., 2003). Although I believe zyxin is being immunolocalized in Figures 3.1-3.4, it is unclear which isoform is being expressed at each specific location.  3.3 Discussion In this study, I demonstrate that zyxin is present at tubulobulbar complexes in the seminiferous epithelium of the testis. Furthermore, zyxin colocalizes with vinculin. Zyxin is a significant component of focal adhesions (Crawford and Beckerle, 1991). The protein contains a number of dynamic structural features including three C-terminal LIM domains involved in proteinprotein interactions, a proline rich domain that can interact with Ena/VASP family members and a nuclear export signal that allows translocation between the nucleus and cytoplasm (Drees et al., 2000; Nix and Beckerle, 1997; Schmeichel and Beckerle, 1994). Zyxin null 3T3 cells display increased motility in comparison to wildtype cells that express zyxin. This phenotype is due to weaker stress fibers that result from a lack of zyxin expression (Hoffman et al., 2003). Re-expression of zyxin suppresses the migratory phenotype (Amsellem et al., 2005). A number of experiments have shown that zyxin distribution in a cell is mechanosensitive. In mouse fibroblasts, changes in the actin cytoskeleton were observed upon the application of uniaxial cyclic stretch. Filamentous actin realigned itself to be perpendicular to the stretch vector (Yoshigi et al., 2005). Zyxin mobilized from  	
    53	
    focal adhesions to stress fibers when cells were subject to the stretch assay. At stress fibers, zyxin acts to reinforce the actin cytoskeleton in response to mechanical stimulation. At focal adhesions in mouse fibroblasts zyxin and vinculin are colocalized. During cell stretching assays, zyxin translocates from the focal adhesion to stress fibers; however, vinculin stays at the focal adhesion (Yoshigi et al., 2005). This is in contrast to what we have observed at tubulobulbar complexes and what Gimona and coworkers (Gimona et al., 2003) observed at smooth muscle podosomes where zyxin and vinculin both relocate from sites of adhesion to the actin cytoskeleton (Kaverina et al., 2003). Tubulobulbar complexes are testis-specific structures that I propose engage in a sort of clathrin-mediated bulk endocytosis of intercellular junctions during sperm release and translocation of spermatocytes. “Clathrin-mediated bulk endocytosis” is a description recently coined in reference to the behavior of reconstituted membranes in a cell free system (Wu et al., 2010). Upon fission disruption with a G-protein analog, clathrin-mediated tubulation was observed resulting in structures that morphologically resemble tubulobulbar complexes (Wu et al., 2010). Tubulobulbar complexes that develop at junctions between Sertoli cells and spermatids consist of a long tubular protrusion of spermatid plasma membrane into a corresponding invagination of the adjacent Sertoli cell (LD, 1979; Russell, 1979a; Russell and Clermont, 1976a). A network of actin filaments  	
    54	
    cuffs the double-membrane tube and the entire structure ends in a clathrincoated pit (Young et al., 2009b). Tubulobulbar  complexes  have  features  in  common  with  podosomes. Podosomes are subcellular actin-containing structures formed at cell/matrix contacts (Linder and Aepfelbacher, 2003). Although the function of podosomes is not entirely clear, adhesion and matrix degradation are likely (Buccione et al., 2004). The formation of podosomes is dependent on an actin assembly complex involving N-WASP, Arp 2/3, cortactin and dynamin (Buccione et al., 2004). During this formation, zyxin and vinculin redistribute from focal adhesions to podosomes (Kaverina et al., 2003). A similar sort of redistribution appears to occur during the formation of tubulobulbar complexes in regions of intercellular attachment in the seminiferous epithelium. In apical regions of the seminiferous epithelium, tubulobulbar complexes form in regions where unique actin-related intercellular adherens-like junctions (ectoplasmic specializations) occur at sites of Sertoli cell attachment to spermatids. Vinculin and zyxin have both been reported to be present at these actin-related junctions (Grove and Vogl, 1989; Lee et al., 2004). The actin assembly proteins N-WASP, Arp2/3 cortactin and dynamin 3 are present at tubulobulbar complexes and presumably are involved with their formation. Vinculin and zyxin appear to redistribute from ectoplasmic specializations to tubulobulbar complexes as the former structures disassemble. The exact role of zyxin and vinculin at tubulobulbar complexes remains to be identified.  	
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    3.4 Chapter 3 figures  Figure 3.1 Fluorescence micrographs of rat seminiferous epithelium triple labeled for actin (phalloidin), zyxin and DNA (DAPI). (A) Stage V of spermatogenesis. The phalloidin (green) stains actin filament bundles in ectoplasmic specializations near the base of the epithelium and at sites of attachment between Sertoli cells and spermatids. Zyxin (red) displays a similar staining pattern localizing to ectoplamic specializations and at sites of attachment between Sertoli cells and spermatids. (B) Stage VII of spermatogenesis. Phalloidin stains tubulobulbar complexes in a similar pattern as zyxin. Zyxin is redistributed from ectoplamic specializations to tubulobulbar complexes in stage VII of spermatogenesis. Arrows indicate location of basal ES. Arrowheads indicate location of apical ES. Bar = 20 mm.  	
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    Figure 3.2 Phase and immunofluorescence micrographs of a rat spermatid and attached sertoli cell regions labeled for zyxin (red), actin (green), and DNA (blue). Actin can be seen outlining individual tubulobulbar complexes that are adjacent to the concave surface of the spermatid head. Zyxin displays a more diffuse, although specific staining around tubulobulbar complexes. Bar = 5 mm  	
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    Figure 3.3 Paired phase and immunofluorescence micrographs of rat spermatids and attached Sertoli cell regions triple labeled for zyxin, vinculin and dapi (blue). (A and B) Zyxin and vinculin display nearly identical staining patterns— confirmation that the two proteins co-localize. (E) Western blot of whole rat testis (WT) and seminiferous epithelium (SE) labeled for vinculin. A single band in each lysate can be observed at 117 kDa, the appropriate molecular weight of vinculin. Bar = 5 mm.  	
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    Figure 3.4 Controls for zyxin labeling of rat seminiferous epithelial fragments. (A) Late spermatid and attached Sertoli cell cuff labeled for zyxin. Diffuse staining is present around tubulobulbar complexes. (B) Late spermatid and attached Sertoli cell treated with Normal Rabbit Serum (NRS) instead of primary antibody. (C) Secondary antibody control. (D) Blank control for autofluorescence. (E) Western blot of seminiferous epithelium and whole testis lysates probed with Zyxin (B71) antibody. A number of immunoreactive bands are visible. The intense band at 20 kDa (asterisk) indicates the HED-2 zyxin splice variant. The 82 kDa band is the expected molecular weight of full length zyxin (arrowhead). Bar = 5 mm.  	
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    Chapter 4 – Tubulobulbar complexes are intercellular podosome-like structures that internalize intact intercellular junctions during epithelial remodeling events in testis.  4.1 Brief synopsis  The current working hypothesis for the function of tubulobulbar complexes is that these structures are novel subcellular machines responsible for internalizing large areas of intact intercellular junctions during sperm release and during translocation of the next generation of developing sperm cells through basal junction complexes. To further characterize tubulobulbar complexes in this study, the following was proposed. If tubulobulbar complexes are responsible for internalizing intact intercellular junctions, then junction molecules should be concentrated at tubulobulbar complexes. A number of adhesion molecules are known to be present in the Sertoli cell plasma membrane at ectoplasmic specializations. Among these molecules are Nectin 2 and α6β1 integrin among others (Gliki et al., 2004; Ozaki-Kuroda et al., 2002; Palombi et al., 1992; Salanova et al., 1995). Also reported to be present is N-cadherin (Yan and Cheng, 2005). The binding partner for nectin 2 is nectin 3 in the spermatid membrane at apical sites and another nectin 2 in adjacent Sertoli cell plasma membrane at basal sites (Ozaki-Kuroda et al., 2002). At apical and basal sites, the ligand for α6β1 integrin is reported to be γ3 laminin  	
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    (Siu and Cheng, 2004) and the ligand for N-cadherin is presumably another Ncadherin molecule in the adjacent membrane. We have shown previously that nectin 2 and nectin 3 are present in the vicinity of tubulobulbar complexes and others have detected what appear to be tight and gap junctions in tubulobulbar complexes based on ultrastructure (Guttman et al., 2004b) (Pelletier, 1988; Russell, 1979c). When tubulobulbar complexes do not form, there is a delay in sperm release – an observation consistent with the junction internalization hypothesis (Kusumi et al., 2007). In this study, the prediction that junction molecules are internalized by and concentrated in tubulobulbar complexes is explored by probing testis sections and fragments with molecules reported to be present at ectoplasmic specializations (nectin 2, nectin 3, α6β1 integrin and N-cadherin) and evaluating the staining at high resolution. Tubulobulbar complexes and related structures were also probed with cortactin – a key podosome component and early endosome antigen 1 (EEA1), a marker for early endosomes. Apical tubulobulbar complexes were the focus of the study because they are most easily visualized at this location. N-cadherin and α6 integrin were labeled in rat and the nectin antibodies used in this study were labeled in mouse because they are not reactive in rat. As predicted, adhesion molecules known to be present in ectoplasmic specializations also are present in tubulobulbar complexes that contain cortactin, a key component of podosomes. Previous findings are extended to show that  	
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    nectin 2 and nectin 3 occur in the elongate tubular parts of the complexes and appear concentrated in vesicular elements at their ends. These vesicles are related to EEA1, indicating that the vesicles are being internalized. Integrins are dramatically concentrated at the ends of tubulobulbar complexes. Surprisingly, NCadherin is not localized to ectoplasmic specializations, nor is it present in tubulobulbar complexes. Rather, N-cadherin is detected only in basal regions of the epithelium and is localized primarily to desmosomes between adjacent Sertoli cells and between early spermatogenic cells and Sertoli cells.  4.2 Results  4.2.1 Nectin 2 Antibodies to the Sertoli cell adhesion molecule nectin 2 reacted both with apical ectoplasmic specializations and with tubulobulbar complexes. This was particularly evident in epithelial fragments double labeled for actin and nectin 2 (Fig. 4.1a). Actin bundles clearly defined ectoplasmic specializations in Sertoli cell regions adjacent to the spermatid head, and these regions also were positive for nectin 2 (Fig. 4.2). Staining of actin filaments associated with each tubulobulbar complex appeared as a “cocoon” or cuff around the tubular region of the complex. These regions also reacted strongly with antibodies to cortactin (Fig. 4.1b) The tubulobulbar complexes were concentrated in the small concavity between the tip and body of the spermatid head. Although nectin 2 staining was  	
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    detected in tubular regions of the complexes, it was most pronounced in vesicular clusters associated with the distal ends of the complexes (Figs. 4.1, a and b, and 4.2a). Vesicles labeled for nectin 2 also were surrounded by a cuff of EEA1 staining (Fig. 4.1c). Similar staining patterns for nectin 2 (Fig. 4.2) were absent in fragments treated with normal rat IgG used at the same concentration as the primary antibody, fragments in which the primary antibody is replaced with buffer alone (secondary antibody control), and fragments in which both primary and secondary antibodies were replaced with buffer (control for autofluorescence).  4.2.2 Nectin 3 Antibodies to the spermatogenic cell adhesion molecule nectin 3 reacted in a similar pattern to antibodies raised against the Sertoli cell adhesion molecule nectin 2 in epithelial fragments of mouse epithelium; that is, the probe labeled ectoplasmic specializations and tubulobulbar complexes (Figs. 4.3 and 4.4). As with the nectin 2 antibody, nectin 3 was concentrated in vesicular clusters associated with the ends of actin-labeled (Fig. 4.3a) and cortactin-labeled (Fig. 4.3b) tubulobulbar complexes, although tubular regions of the complexes also were stained (Fig. 4.4). Nectin 3 labeled vesicles at the ends of tubulobulbar complexes also labeled for EEA1 (Fig. 4.3c). A similar pattern present for nectin 3 staining was not present in controls (Fig. 4.4, e-g).  	
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    4.2.3 α6 integrin Labeling with antibody generated against α6 integrin was intense and specifically associated with the ends of tubulobulbar complexes and related vesicles (Figs. 4.5 and 4.6). Using our staining protocols, the antibody was not reactive above background with α6 integrin in ectoplasmic specializations. Controls for staining were negative (Fig. 4.6). On Western blots of testis and seminiferous epithelium (Fig. 4.6e), the antibody labeled four bands not present in IgG controls – one at the appropriate molecular weight for full-length α6 integrin (120 kDa) and three at lower molecular weights. The band with the lowest molecular weight (around 40 kDa) was the most reactive.  4.2.4 N-Cadherin is concentrated at desmosome junctions and not at ectoplasmic specializations or tubulobulbulbar complexes The antibody to N-cadherin labeled intercellular contacts in basal regions of the seminiferous epithelium at all stages of spermatogenesis, but was not reactive with apical ectoplasmic specializations associated with spermatids at any stages of spermatogenesis, nor was it reactive with tubulobulbar complexes associated with late spermatids (Fig. 4.7). Staining controls were negative (Fig. 4.7) and the antibody reacted with a single band at the appropriate molecular weight (135 kDa) for N-cadherin on Western blots of testis and seminiferous epithelium (Fig. 4.7).  	
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    The absence of N-cadherin staining in apical regions was evident in frozen sections of fixed testis single-labeled for N-cadherin (Fig. 4.7), and was even more clearly evidenced in sections double labeled for N-cadherin and for Factin (to indicate the locations of apical ectoplasmic specializations and tubulobulbar complexes; Fig. 4.8) No staining for N-cadherin was detectable at these sites. In basal regions of the epithelium, labeling with the N-cadherin antibody did overlap to some extent with the location of actin bundle staining. However, this overlap was not absolute and was stage specific; that is, there was more overlap at some stages (for example, stage V) than at others (for example, stages X and VII; Fig. 4.8). Moreover, staining with the N-cadherin often appeared punctate and occurred in regions where there were no actin filament bundles. To explore further the location of N-cadherin at junction sites, thin sections of rat testis were probed with the antibody at the ultrastructural level using immunogold techniques (Fig. 4.9). As expected on the basis of immunofluorescence results, the antibody was not reactive with apical ectoplasmic specializations (Fig. 4.9a). At basal junction complexes between adjacent Sertoli cells, gold particles were clustered at focal locations and were not distributed uniformly along regions of the complexes containing ectoplasmic specializations (Fig. 4.9 b and c). In addition, gold particles occurred at sites of contact between early spermatogenic cells and Sertoli cells that clearly were not associated with ectoplasmic specializations based on location and on the  	
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    absence of any actin filament bundles. The reactive sites in regions of contact between neighboring Sertoli cells and between Sertoli and spermatogenic cells were clearly related to intermediate filaments (Fig. 4.9, c and e-g). To explore further the relationship at the ultrastructural level between Ncadherin staining and junctions, sections were double labeled for N-cadherin and for beta actin. Consistent with previous results, N-cadherin was concentrated at desmosomes and not at actin filament-rich apical and basal ectoplasmic specializations (Fig. 4.10). Significantly, some actin staining above background also was detected at desmosomes and appeared closely related to the submembrane plaques (Fig. 4.10 d-f). To summarize, N-cadherin is localized to desmosomes in the seminiferous epithelium and is not concentrated at ectoplasmic specializations or at tubulobulbar complexes.  4.3 Discussion In this study, integral membrane junction molecules were demonstrated to be present at apical adhesion junctions between Sertoli cells and spermatids, and also are present in podosome-like tubulobulbar complexes and appear concentrated and are internalized at their ends. The results from this study support the general hypothesis that tubulobulbar complexes internalize intact intercellular junctions during junction disassembly. Pertinent to the disassembly hypothesis are a number of key observations. First, tubulobulbar complexes develop at intercellular junctions,  	
    66	
    indicating that there is a functional relationship between tubulobulbar complexes and junctions (Russell and Clermont, 1976a). Significantly, tubulobulbar complexes occur not only at apical junctions between Sertoli cells and spermatids, but also at basal junctions between adjacent Sertoli cells, indicating that the primary function of tubulobulbar complexes is not related just to spermatid maturation (Russell, 1979a; Vaid et al., 2007). Second, junction elements are present in tubulobulbar complexes. Ultrastructurally, identifiable gap and tight junctions have been reported in basal junction complexes, and this study found that adhesion molecules (nectin 2, nectin 3, and α6 integrin) known to be present at apical junctions between Sertoli cells and spermatids, also are present in apical tubulobulbar complexes and appear concentrated at their ends. The finding that nectin 3 is concentrated in clusters of vesicles at the ends of tubulobulbar complexes is particularly significant because this adhesion molecule is present in the spermatid plasma membrane at apical ectoplasmic specializations and yet appears to be associated with vesicles being internalized into Sertoli cells. This observation, together with the findings that nectin 2 in the Sertoli cell plasma membrane has a similar distribution and that these nectin-2 and nectin-3 labeled vesicles also stain with antibodies to EEA1, supports the conclusion that intact junctions are being internalized at tubulobulbar complexes. The finding that α6 integrin is localized to tubulobulbar complexes is consistent with the staining of the β1 integrin subunit at tubulobulbar complexes in images of seminiferous epithelium  	
    67	
    presented in the original report of β1 integrin in the testis (Palombi et al., 1992). In a subsequent study from the Vogl lab, β1 integrin was specifically not reported at tubulobulbar complexes. This is possibly because the antibody used was different than that used in the original study and did not stain the structures. Unfixed material was used in this study so the morphology was poor and tubulobulbar complexes were not well preserved, also, the material was observed at low magnification because tubulobulbar complexes were not the focus of the study therefore, the staining went unnoticed. Significantly, the antibody to the α6 integrin subunit (a polyclonal antibody raised against residues 868-954 of human α6 integrin near the C-terminus of the protein) is not highly reactive with rat ectoplasmic specializations in tissue processed using the fixation techniques used in this study, but intensely labels the ends of tubulobulbar complexes and associated clusters of vesicles. It is possible that the epitope on this integrin subunit may be masked in ectoplasmic specializations but becomes exposed as the integrin is moved into tubulobulbar complexes and parts of the extracellular domain are proteolytically cleaved. Supporting this interpretation of the data is the observation on Western blots of testis and seminiferous epithelium; the antibody reacts strongly with a band that migrates at a lower molecular weight than that of the full-length intact heavy chain of the α6-integrin subunit. This band is not present in control lysates from INT407 cells, and it may be similar to that described in certain invasive and metastatic cancer cells, where the β-barrel extracellular domain of α6 integrin is cleaved by plasminogen activator  	
    68	
    (Demetriou and Cress, 2004; Demetriou et al., 2004; Pawar et al., 2007). The plasminogen activation system has been the focus of intense study in the testis (Le Magueresse-Battistoni, 2007). The results from this study may provide a molecular link between this system and a normal physiological event (junction turnover) through cleavage of the α6 integrin subunit. However, there is at least one caveat to the interpretation of the Western blot data. The testis and seminiferous epithelial lysates were prepared in the absence of protease inhibitors, whereas the INT 407cells were prepared in the presence of protease inhibitors. It is possible that the staining intensity of the lower-molecular weight band was exaggerated in our blots of testis and seminiferous epithelium and that the intensity of the full-length α6 integrin subunit band was weakened. Third, clusters of double-membrane-bound vesicles are observed by electron microscopy at the ends of tubulobulbar complexes – an observation consistent with the proposal that bulbar regions of the complexes bud from tubulobulbar complexes and “intact” junctions are internalized by Sertoli cells (Guttman et al., 2004b). Fourth, vesicles associated with tubulobulbar complexes have been shown previously to label positively for lysosomal and endosomal markers, indicating that tubulobulbar complexes are related in some way to internalization and degradation pathways (Guttman et al., 2004b; Russell and Clermont, 1976a; Russell, 1979c).  	
    69	
    Fifth, when tubulobulbar complexes do not form, there is reportedly a delay in sperm release. This result would be predicted if tubulobulbar complexes played a role in normal junction disassembly (Kusumi et al., 2007). Although during junction turnover in other systems, integral membrane junction molecules in adjacent plasma membranes disengage and are internalized into each of their parent cells by conventional endocytic mechanisms the model proposed in this thesis for junction internalization by tubulobulbar complexes in the seminiferous epithelium is not without precedent. Generally, in cells, gap junctions, consisting of connexions in the plasma membranes of two attached cells are internalized into one or the other of the attached cells as intact junctions in double- membrane vesicles and the process is clathrin dependent (Berthoud et al., 2004; Gaietta et al., 2002; Piehl et al., 2007). A similar type of phenomenon occurs with tight junctions in certain situations (Matsuda et al., 2004). In Sertoli cells, tubulobulbar complexes may be the mechanism by which massive intact intercellular junctions can be internalized at specific sites while still maintaining some form of connection between cells. This mechanism of internalization also would facilitate Sertoli cells removing from the surfaces of maturing spermatids junction molecules that would no longer be required by these cells following sperm release. The finding that N-Cadherin is primarily concentrated in desmosomes between adjacent Sertoli cells and between Sertoli cells and early spermatogenic cells is generally consistent with some reports on the localization of cadherins in  	
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    the seminiferous epithelium, but it is inconsistent with others in which N-cadherin is reported to be a major component of ectoplasmic specializations both at apical and basal junctions (Johnson and Boekelheide, 2002; Mulholland et al., 2001; Yan and Cheng, 2005). Part of the discrepancy may be due to different processing techniques and to the fact that at basal sites, desmosomes between adjacent Sertoli cells are intercalated into basal junction complexes, making it difficult to differentiate between ectoplasmic specializations and other junction types. This study is the first to evaluate N-cadherin staining at the ultrastructural level. Based on our results, we propose that desmosomes in the seminiferous epithelium may be a hybrid form of junction related both to the actin cytoskeleton and to intermediate filaments, and containing conventional in addition to possibly desmosomal cadherins. The observation that some actin is detectable above background is consistent with this possibility. The selection pressure for the evolution of this type of junction may be related to the lack of cytoplasmic intermediate filament expression in spermatogenic cells. A hybrid junction would allow cortical actin filaments to associate with the junction on the spermatogenic cell side of the junction, whereas on the Sertoli cell side the junction could be related both to intermediate and perhaps also actin filaments. The lack of Ncadherin staining in apical tubulobulbar complexes is consistent with the lack of staining in the related ectoplasmic specializations.  	
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    4.4 Chapter 3 figures  	
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    Figure 4.1 Phase and immunofluorescence micrographs of mouse spermatids and attached sertoli cell regions triple-labeled (a) actin, nectin 2, and DNA (DAPI); (b) cortactin nectin 2 and DNA (DAPI); and (c) EEA1, nectin 2, and DNA (DAPI). In a, the actin stain clearly labels the actin networks of tubulobulbar complexes, as well as the actin bundles in ectoplasmic specializations (ES) in Sertoli cell regions overlying the spermatid heads. The nectin 2 antibody labels tubulobulbar complexes and appears concentrated near their ends (arrowhead). The antibody also labels the plasma membrane of Sertoli cells at ectoplasmic specializations (ES). In b, the antibody to cortactin strongly labels tubulobulbar complexes, and nectin 2 is concentrated in structures (arrowhead) associated with their ends. In c, EEA1 (arrowhead) is associated with the vesicular structure that stains intensely for nectin 2. Bar = 1 μm.  	
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    Figure 4.2 Paired phase and immunofluoresence micrographs of mouse epithelial fragments single labeled for nectin 2. a) Spermatid head and related Sertoli cell regions single labeled for nectin 2. Nectin 2 staining is associated with tubulobulbar complexes (arrowheads). b) Epithelial fragment treated with normal rat IgG (N Rat IgG). c) Secondary antibody control (2° Control). d) Control for auto-fluorescence (Blank). Bars = 1 μm.  	
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    Figure 4.3 Phase and immunofluorescence micrographs of mouse spermatids and attached sertoli cell regions triple labeled (a) actin, nectin 3, and DNA (DAPI); (b) cortactin, nectin 3, and DNA (DAPI); and (c) EEA1, nectin 3, and DNA (DAPI). In a, the actin stain outlines tubulobulbar complexes and also labels ectoplasmic specializations (ES). The nectin 3 	
    75	
    antibody appears mainly concentrated near the ends of the tubulobulbar complexes (arrowhead). The antibody also labels regions (asterisk) related to the attached ectoplasmic specializations of the adjacent Sertoli cell. In b, the cortactin antibody intensely labels tubulobulbar complexes, and nectin 3-positive structures (arrowhead) are associated with their ends. In c, EEA1 staining (arrowhead) is associated with a nectin 3-positive vesicle (arrowhead). Bar = 1 μm.  	
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    Figure 4.4 Paired phase and immunofluorescence micrographs of mouse epithelial fragments containing late spermatids and adjacent sertoli cell regions single labeled for nectin 3. a–d) Epithelial fragments labeled for nectin 3. Labeling clearly appears concentrated at the ends of tubulobulbar complexes (arrowheads), but also can be detected in the elongate tubular regions of the complexes (arrows). The antibody also reacts with the plasma membrane of the spermatid heads (asterisks) in regions adjacent to ectoplasmic specializations of Sertoli cells. e) Epithelial fragment treated with normal rat IgG (N Rat IgG) rather than primary antibody. f) Secondary antibody control (2° Control). g) Control for autofluorescence (Blank). Bars = 1 μm.  	
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    Figure 4.5 Epithelial fragments of rat seminiferous epithelium triple labeled for actin, α6 integrin, and DNA (DAPI). a and b) Tubulobulbar complexes and ectoplasmic specializations (ES) are labeled with the probe for filamentous actin. The α6 integrin antibody labels the ends of individual tubulobulbar complexes (small arrows) and large masses associated with them (arrowheads). Bar = 5 μm.  	
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    Figure 4.6 Controls for α6 labeling of rat seminiferous epithelial fragments. a) Late spermatid and adjacent Sertoli cell regions labeled for α6 integrin. Staining is concentrated at the ends of tubulobulbar complexes (arrowhead). b) Apical epithelial fragment treated with normal rabbit IgG (NRIgG) instead of primary antibody. c) Secondary antibody control. d) Control for autofluorescence. e) Western blot of rat testis and seminiferous epithelium (SE) labeled for α6 integrin. Notice the upper band (arrowhead) that is at the molecular weight (kDa) expected for the heavy chain of α6 integrin is almost invisible in whole testis and is moderately stained in seminiferous epithelium and strongly stained in INT407 cells. Also notice the strong lower-molecular weight band (asterisk) in testis and particularly in seminiferous epithelium, but not in INT407 cells, that migrates just below 43 kDa. Bars = 5 μm.  	
    79	
    Figure 4.7 Paired phase and immunofluorescence micrographs of fixed frozen sections of rat testis single labeled for N-cadherin. a) Section of seminiferous epithelium at stage V of spermatogenesis. Labeling is confined to basal regions of the epithelium (white arrow) and is not associated with apical regions surrounding spermatid heads. b) Section of seminiferous epithelium at stage VII of spermatogenesis. Labeling occurs only at the base of the epithelium (white arrows) and does not label ectoplasmic specializations or tubulobulbar complexes associated with late spermatids near the apex of the epithelium. c) Section of epithelium at stage VII of spermatogenesis treated with normal mouse IgG (NMIgG) instead of primary antibody. d) Control for secondary antibody (2° Control). e) Control for autofluorescence (Blank). Bars = 10 μm. f) Western blot of rat testis and seminiferous epithelium labeled for N-cadherin. The antibody is monospecific for a protein approximately around 135-kDa molecular mass (asterisk-labeled arrow).  	
    80	
    Figure 4.8 Phase and fluorescence micrographs of rat seminiferous epithelium triple labeled for actin, n-cadherin, and DNA (DAPI). a) Stage X of spermatogenesis. b) Stage V. c) Stage VII. The actin stain labels filament bundles in ectoplasmic specializations at the base of the epithelium (small white arrows) and at sites of attachment between Sertoli cells and spermatids (arrows indicated by the asterisks). At stage VII, tubulobulbar complexes are also labeled with the actin probe. The N-cadherin antibody only labels structures near the base of the epithelium and does not label apical ectoplasmic specializations or tubulobulbar complexes. Some of these features overlap with the actin staining, particularly at stage V (b), whereas others (small white arrowheads) clearly are not colocalized with actin. Bars = 10 μm.  	
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    Figure 4.9 Immunoelectron microscopic images of rat testis sections labeled for n-cadherin. a) Spermatid head and associated ectoplasmic specialization (ES) in the surrounding Sertoli cell. No gold particles occur in the region of the ectoplasmic specialization. Some nonspecific staining occurs over the nucleus of the spermatid head. b) Basal junction complex between two adjacent Sertoli cells. Gold particles (arrowheads) occur in regions of the complex associated with intermediate filaments (IFs), but do not occur in regions associated with ectoplasmic specializations (ES). c) Cluster of gold particles (arrowhead) in a region of a basal junction complex associated with intermediate filaments (IFs). d) Section of a desmosome between a Sertoli cell and a round germ cell treated with normal mouse IgG instead of the primary antibody. Notice that intermediate filaments (IFs) occur only on the Sertoli cell side of the junction and that no gold particles are present at the plasma membranes. e and f) Sections of desmosomes between Sertoli cells and round germ cells labeled for N-cadherin. Gold particles are associated with the membranes of both cells at the junctions. Notice the intermediate filaments (IFs) on the Sertoli cell side of the desmosomes. g) Section of a desmosome intercalated into the basal junction complex between two neighboring Sertoli cells. Gold particles are present at the intermediate filament (IF)-associated desmosome, but are absent from the area occupied by ectoplasmic specializations (ES). Bars = 100 nm.  	
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    Figure 4.10 Immunoelectron microscopic images of rat testis double labeled for n-cadherin and actin. Ectoplasmic specializations (ES) both at basal sites of attachment to adjacent Sertoli cells (a) and apical sites of attachment to spermatids (b) label only with the probe for actin (5-nm gold particles) and do not label with the antibody to Ncadherin (10-nm gold particles). When the primary antibodies are replaced with normal mouse IgG (NMIgG) and normal rabbit IgG (NRIgG), no labeling above background occurs at ectoplasmic specializations. c) The antibody for N-cadherin (black arrowheads) clearly labels desmosomes at basal sites of attachment between Sertoli cells and germ cells (d–f). Interestingly, the probe for actin 	
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    (arrows) also occurs above background levels at desmosomes and labels material close to the plasma membrane. No specific labeling is present when the primary antibodies are replaced with normal mouse and normal rabbit IgG. Bar = 100 nm.  	
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    Chapter 5 – Internalization of adhesion junction proteins and their association with recycling endosome marker proteins in rat seminiferous epithelium. 5.1 Brief synopsis In the previous chapter, the adhesion molecules nectin 2, nectin 3 and α6 integrin were localized to tubulobulbar complexes. Furthermore, nectin 2 and nectin 3 were localized in the vesicles associated with tubulobulbar complexes that labeled positively for EEA1. Previous to this study, other adhesion proteins and endosomal markers had not yet been localized to these sites in rat. In this study, β1 integrin and nectin 2 are found to be concentrated at the ends of tubulobulbar complexes in the rat, and for the first time in any species it is demonstrated that these regions are also associated with the early endosomal marker Rab5. Also, the ʻlong-loopʼ recycling pathway marker Rab11 is shown to occur at vesicular-like structures where adhesion junctions are newly forming adjacent to early spermatids and the marker (Caswell and Norman, 2006). The Rab11 marker is co-distributed with nectin2 at these sites. ʻShort-loopʼ and ʻlongloopʼ endocytic pathways refer to two spatially and temporally distinct mechanisms controlled by Rab4 and Rab11 GTPases respectively. The codistribution of nectin 2 and Rab11 is consistent with the hypothesis that tubulobulbar complexes at the apical sites of attachment internalize adhesion junctions in the rat. Furthermore, the results from this study indicate for the first  	
    86	
    time that at least some of the junction proteins may be recycled to newly forming junctions with the next generation of spermatids.  5.2 Results  5.2.1 Nectin 2 and the β1 integrin subunit are concentrated at the ends of apical tubulobulbar complexes Antibodies to both nectin 2 (Fig. 5.1) and β1 integrin (Fig. 5.2) labeled Sertoli cell regions adjacent to the heads of late spermatids consistent with the position of ESs and material in Sertoli cell apical lobules at the ends of tubulobulbar complexes. The probe for nectin 2 clearly labeled basal junction complexes between Sertoli cells and regions of Sertoli cell attachment to spermatids. This was very evident at stage V of the seminiferous epithelium (Fig. 5.1) where spermatids were entrenched deep within apical Sertoli cell crypts. This staining pattern overlapped with the staining of filamentous actin known to be concentrated in ESs at adhesion junctions. At stage VII, when mature spermatids were at the apex of the epithelium, nectin 2 staining was also co-distributed with actin staining at apical and basal ESs (Fig. 5.1) In addition, focal nectin 2 staining occurred in the central regions of Sertoli cell apical lobules surrounding late spermatids (arrowheads in Fig. 5.1), regions known to contain tubulobulbar complexes. At higher magnification of epithelial fragments labeled for actin, this 	
    87	
    staining was concentrated at the ends of tubulobulbar complexes (Fig. 5.1). Similar staining patterns at ESs and at the ends of tubulobulbar complexes were absent when the primary antibody was replaced with the same concentration of normal rabbit IgG (Fig. 5.1), when the primary antibody was replace with buffer alone (Fig. 5.1) and when both primary and secondary antibodies were replaced with buffer alone (Fig. 5.1). In addition, the antibody reacted with a single band of the appropriate molecular weight for nectin 2 (60-65 kDa) on immunoblots of whole testis and isolated seminiferous epithelium not present on control blots treated with normal rabbit IgG instead of the primary antibody (Fig 5.1). In epithelial fragments processed for antigen retrieval, the antibody to the β1 integrin subunit reacted with two regions associated with late spermatids: regions known to contain ESs adjacent to the dorsal or convex surface of spermatid heads and regions of Sertoli cell lobules known to contain distal parts of tubulobulbar complexes (Fig. 5.2) Staining associated with the dorsal curvature of spermatid heads was intense while that associated with tubulobulbar regions was weaker and more diffuse. Staining with fluorescent phallotoxins for filamentous  actin  to  highlight  tubulobulbar  complexes  and  ectoplasmic  specializations in this material was not successful because glutaraldehyde is used in the fixation protocol. Controls for staining were all negative (Fig. 5.2).  	
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    5.2.2 The early endosomal marker rab5 is distinctly localized at the ends of tubulobulbar complexes. When epithelial fragments containing late spermatids and associated Sertoli cell apical lobules were labeled both for filamentous actin and for Rab5, staining for Rab5 occurred only at the distal ends of tubular regions of tubulobulbar complexes outlined by actin labeling (Fig. 5.3a-d) Small segments of actin staining also appeared to occur distal to the Rab5-stained regions. In material fixed and processed for antigen retrieval, staining at the ends of tubulobulbar complexes was dramatically evident (Fig. 5.3e). The Rab5 staining occurred in regions similar to those reported above for nectin 2 and β1 integrin. Controls for Rab5 staining were all negative (Fig. 5.3f,g,h) and the antibody reacted with a major band of the appropriate molecular weight (24 kDa) for Rab5 on immunoblots of testis and seminiferous epithelial lysates (Fig. 5.3i). Rab5 staining on blots of whole testis and seminiferous epithelium was similar, indicating that tissues other than the epithelium contain significant levels of Rab5.  5.2.3 The ʻlong-loopʼ recycling marker rab11 is localized to newly forming junctions associated with early spermatids Staining with the immunological probe for Rab11 was dramatically evident in epithelial fragments adjacent to spermatids that were in the early stages of elongation (Fig. 5.4) Here, the labeling pattern was vesicular in appearance with the vesicles closely related to ectoplasmic specializations indicated by actin  	
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    staining. In early spermatids that were beginning to polarize (step 8 spermatids), Rab11 positive puncta were often around the margins of the developing ectoplasmic specializations (Fig. 5.4a) whereas somewhat later (step 9/10) the vesicular staining was more widespread over the actin-positive junction (Fig. 5.4b,c). Similar staining patterns were not observed in epithelial fragments treated with normal rabbit IgG at the same concentration as with primary antibody (Fig. 5.4d), when the primary antibody was replaced with buffer alone (Fig. 5.4e), or when both the primary and secondary antibodies were replaced with buffer alone (Fig. 5.4f). On immunoblots of whole testis and isolated seminiferous epithelium, the antibody reacted with a single major band at the appropriate molecular weight (22 kDa) for Rab11 (Fig. 5.4g) and like Rab5, the protein did not appear enriched in seminiferous epithelium.  5.2.4 Rab11 is co-distributed with nectin 2 at newly forming junctions To determine whether the junction molecules were co-distributed with Rab11, epithelial fragments were double labeled with probes for Rab11 and nectin 2. Significantly, no Rab11 staining occurred in Sertoli cell lobules associated with late spermatids, either at nectin 2-positive ESs or at nectin 2positive vesicles at the ends of tubulobulbar complexes (Fig. 5.5a). However, puncta clearly double labeled for nectin2 and Rab11 were associated with regions known to contain ESs attached to early spermatids (Fig. 5.5b, 5.6a,b). When sections containing step 11 spermatids were labeled for nectin 2 and  	
    90	
    Rab11 and then analyzed using ImageJ (National Institutes of Health, Bethesda, MD, USA) with a co-localization plug-in (Abramoff et al., 2004), the probes were co-localized in vesicular structures associated with apical junctions (Fig. 5.7) When additional sections were analyzed with Image J, it was confirmed that co-localization of nectin 2 and Rab11 was stage specific. There was little colocalization at apical junction sites in stages V and VII (Fig. 5.8a,b), whereas in stages containing early elongating spermatids where apical adhesion junctions (ie. ESs) were newly forming (Fig. 5.8c), there was significantly more colocalization. Interestingly, co-localization at basal junctions between Sertoli cells was also more apparent (arrowheads in Fig. 5.8c) in stages containing early elongated spermatids. Here, basal junctions are turning over to allow the translocation of spermatocytes from basal to adluminal compartments of the epithelium.  5.3 Discussion In this study, both nectin 2 and β1 integrin are shown to be present at the ends of tubulobulbar complexes in rat seminiferous epithelium and that similar regions are also associated with early endosomal marker Rab5. Also, nectin 2 and long-loop recycling marker Rab11 co-distribute at vesicles associated with ESs forming with early spermatids. These data presented here are consistent with the conclusion that tubulobulbar complexes internalize intercellular junctions during junction  	
    91	
    remodeling in the seminiferous epithelium. Significantly, Rab5 clearly labels the ends of tubulobulbar complexes in areas that appear proximal to those labeled by EEA1 in mouse (previous chapter). Generally in the endocytosis pathway, Rab5 acts upstream of EEA1 (Christoforidis et al., 1999). The Rab5-positive regions of tubulobulbar complexes may be the bulbar regions because they do not label for actin and are situated precisely at the ends of strongly actin-labeled tubular regions. The concentration of adhesion molecules at the ends of tubulobulbar complexes and the presence of early endosomal markers in similar locations strongly  support  the  argument  that  tubulobulbar  complexes  internalize  intercellular junctions. The fates of junction proteins internalized by tubulobulbar complexes have not been studied; however, by analogy with other systems, the proteins should be degraded and/or recycled. The appearance of what appear to be late endosomes/lysosomes in vesicular clusters associated with the ends of tubulobulbar complexes indicates that at least some of this material is degraded (Guttman et al., 2004b; Russell, 1979b). Also, nectin 3 in spermatid plasma membranes that is internalized by the Sertoli cell would presumably be degraded because this protein is not expressed by Sertoli cells. The situation may be different for nectin 2 and α6β1 integrin. Both are expressed by Sertoli cells and are concentrated at ESs. Significantly, massive amounts of each presumably are internalized during sperm release at the apex of the epithelium while at the same time being added to the plasma membrane deeper in the epithelium in  	
    92	
    association with newly forming junctions with the with the next generation of spermatids. Rab11 regulates trafficking into and out of the recycling endosome (Tarbutton et al., 2005). It has been implicated in the recycling of membrane components and transcytosis of molecules in polarized epithelia (Casanova et al., 1999; Xu et al., 2011). The observation that nectin 2 positive vesicles that also label for Rab11 occur at newly forming ESs with early spermatids is the first indication that junction proteins internalized at tubulobulbar complexes may be recycled to newly forming junctions elsewhere in the epithelium (Fig. 5.9) Based on the tendency of tubulobulbar complexes to occur in association with basal junction complexes between Sertoli cells at the sites of the ʻbloodtestisʼ barrier, junctions may be internalized by tubulobulbar complexes during basal junction turnover when spermatocytes translocate from basal to adluminal compartments of the epithelium. However, except for a report of tight and gap junctions being visible in electron micrographs of tubulobulbar complexes, evidence for this mechanism is lacking (Russell, 1979c). During translocation, junctions disassemble above spermatocytes and new junctions assemble below (Fig. 5.9) (Russell, 1977b). There is evidence from cultured Sertoli cells that junction proteins are endocytosed and recycled back to intercellular junctions, however, it is not known whether this process involves tubulobulbar complexes or how these results translate into the intact seminiferous epithelium. One can speculate that recycling of junction proteins from tubulobulbar complexes may occur at basal junctions in a fashion similar to what occurs at apical sites. The  	
    93	
    observation that nectin 2 and Rab11 are significantly more co-distributed at basal junctions at the stages of spermatogenesis where basal junctions are turning over to allow the next generation of spermatocytes through basal junction complexes is consistent with this argument. If recycling from basal tubulobulbar complexes does occur, it would be interesting to know whether adhesion proteins from apical and basal sites enter the same recycling compartments or are segregated. It also raises interesting questions about how the different junction proteins (adhesion, gap, and tight), which are presumably internalized together by tubulobulbar complexes at basal sites, are each trafficked through membrane recycling pathways. Another issue yet to be clarified is whether junction proteins are differentially  segregated  by  tubulobulbar  complexes  themselves  during  internalization. For example, nectin 2 and nectin 3 might be entirely removed from apical junctions by tubulobulbar complexes, whereas α6β1 integrin may be only partially removed. This may account for the presence of an integrin-based disengagement complex remaining as the last area of contact between Sertoli cells and spermatids after ESs and tubulobulbar complexes have disappeared (Beardsley and O'Donnell, 2003; Beardsley et al., 2006). In this chapter, early endosome marker Rab5 was localized to the ends of tubulobulbar complexes at apical sites of attachment between Sertoli cells and late spermatids in rat. Moreover, β1 integrin and nectin 2 also are concentrated at these sites. Together, these data are consistent with the hypothesis that  	
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    tubulobulbar complexes internalize apical junctions in the seminiferous epithelium. The results from this study also demonstrate that Rab11 and nectin 2 are co-localized at newly forming junctions with the next generation of spermatids and raise the interesting possibility that some of the junction proteins internalized during spermiation may be recycled to junctions forming deeper in the epithelium.  	
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    5.4 Chapter 5 figures  Figure 5.1 Grouped phase and fluorescence images showing the distribution of nectin 2 and actin in rat seminiferous epithelium. Shown in (a) and (b) are the distribution of nectin 2 and actin at stage V and stage VII respectively of the seminiferous epithelium. At both stages, nectin 2 is concentrated at ectoplasmic specializations (ESs), indicated by positive 	
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    phalloidin staining, both at basal junction complexes and at apical sites of attachment to spermatids. At stage VII (b) nectin 2 also is concentrated at focal sites (arrowheads) in apical lobes that envelop late spermatid heads. At higher magnification (c) each of these focal sites appears as a cluster of small vesicularlike structures (arrowhead) at the ends of tubulobulbar complexes identified by actin labeling [inset in (c)]. Similar clusters are absent in material treated with normal rabbit IgG at the same concentration as primary antibody (d), with buffer alone instead of primary antibody (e), and with buffer instead of both the primary and the secondary antibodies (f). In Western blots of rat testis and seminiferous epithelium (g), the nectin 2 antibody reacts strongly with a band (asterisk) between 60-65 kDa that is not present in blots treated with normal rabbit IgG (NRIgG) at the same concentration as primary antibody. Bars = 10 mm (a,b) and 5 mm (c-f)  	
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    Figure 5.2 Paired phase and fluorescence micrographs of late spermatids and associated Sertoli cell regions of the rat processed by the antigen retrieval protocol as indicated in the text and labeled with β1 antibodies. The antibody clearly labels material in apical Sertoli cell lobules in regions known to contain the ends of tubulobulbar complexes [arrowhead in (a)]. Regions known to contain ectoplasmic specializations label intensely with the antibodies [small arrow in (a)]. Similar patterns of staining are not present in material treated with normal rabbit serum at the same concentration as primary antibody (c), buffer instead of primary antibody (d), and buffer instead of both primary and secondary antibodies. Bar = 5 mm  	
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    Figure 5.3 Phase and fluorescence images of late spermatids and associated sertoli lobes mechanically dissociated from perfusion fixed rat seminiferous epithelium and labeled for rab 5. The epithelial fragment containing the spermatid shown in panels (a) to (d) was processed using our conventional immunofluorescence protocol. Tubulobulbar complexes are clearly outlined by the actin labeling in (b). Rab 5 labels regions of the complexes near their ends [arrowheads in (c)]. These regions may correspond to the bulbar regions of the complexes because they occur precisely at the ends of the actin-related proximal tubular regions [arrowheads in (d)]. There also is some actin staining distal to the Rab 5 positive sites [arrows in (d)] in regions that may correspond to the short distal tubular regions of the complexes. Shown in (e) is Rab 5 staining in an epithelial fragment processed 	
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    through the antigen retrieval protocol described in the text. The Rab 5 antibody clearly labels tubulobulbar complexes near their ends (arrowhead). A similar pattern is absent in fragments treated with normal rabbit IgG at the same concentration as primary antibody (f), buffer instead of the primary antibody (g) and buffer instead of both the primary and the secondary antibodies (h). Bars = 5 mm. Panel (i) is a Western blot of rat testis and seminiferous epithelium labeled for Rab 5. The Rab 5 antibody reacts strongly with a major band of the appropriate molecular weight (24 kDa) for Rab 5 on blots both of seminiferous epithelium and testis. Blots were re-probed for actin to control for loading (shown as the lower panel in the figure).  	
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    Figure 5.4 Paired phase and fluorescence micrographs of early spermatids labeled for actin, rab 11 and DNA. Shown in panels (a) and (b), while controls for Rab 11 labeling are shown in panels (c) to (g). The spermatids in (a) and (b) are approximately at steps 8 and 9-10 of spermiogenesis respectively. Positive labeling for Rab 11 occurs in a vesicular-like pattern (arrowheads) around the edge of the developing ectoplasmic specialization indicated by the actin labeling in (a). In (b) the positive vesicles (arrowheads) are more widely distributed over the developing junction. Bar = 5 mm. In panel (c), Rab 11 positive staining occurs in a punctate or vesicular pattern (arrowheads) adjacent to the early spermatid. Similar patterns are not seen in fragments incubated with normal rabbit IgG instead of primary antibody (d), in secondary antibody controls (e) in which the primary antibody is replaced with buffer alone, or when both the primary and secondary antibodies are replaced with buffer alone (f). The Rab 11 antibody reacts strongly with a single band on Western blots of seminiferous epithelium and testis that is not present on similar blots treated with normal rabbit IgG alone (g). Blots were reprobed for actin (lower panel in the (g)) to control for loading. Bars = 5 mm  	
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    Figure 5.5 Phase and fluorescence micrographs of spermatids and associated sertoli cell regions double labeled with antibodies to nectin 2 and rab 11. In association with late spermatids (a), the nectin 2 probe intensely labels ectoplasmic specializations (arrow) associated with the spermatid head and also labels material in regions known to contain the ends of tubulobulbar complexes (arrowhead). Rab 11 staining is negative in association with late spermatids. In association with early elongate spermatids (b) nectin 2 and Rab11 positive vesicles occur in Sertoli cell regions adjacent to the acrosome where adhesion junctions are being assembled. Some of these vesicles appear labeled for both probes. Notice that some of the vesicles are nectin 2 positive and Rab 11 negative. These vesicles may contain newly synthesized protein. Bars = 5 mm  	
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    Figure 5.6 Immunofluorescence micrographs of early spermatids at step 8 (a) and step 9 (b) of spermiogenesis double-labeled for nectin 2 and rab 11. The cells also are stained with DAPI to highlight the nuclei. Both Rab 11 and nectin 2 clearly label the same vesicular or punctate structures (arrowheads) in Sertoli cell regions overlying the acrosome regions of early spermatids. Also notice that some of the vesicles that are Rab 11 positive do not label for nectin 2. These vesicles may contain proteins other than nectin 2 that also are being recycled. Bars = 5 mm  	
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    Figure 5.7 Shown here is a cryo-section of a seminiferous tubule containing approximately at stage XI containing step 11 spermatids. (a) Labeled both for Rab 11 (b) and for nectin 2 (c). The merged image is shown in panel (d). The image was analyzed in ImageJ and regions of co-localization are shown in white in the insets in (d) [arrowheads in (d) indicate the regions enlarged in the insets]. The labeling with both probes is in regions known to contain ectoplasmic specializations. Bar = 20 mm  	
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    Figure 5.8 Cryo-sections of seminiferous tubules at stages V, VII and IX Shown here are cryo-sections of seminiferous tubules at stages V (a), VII (b) and IX (c) of spermatogenesis that have been stained for nectin 2 and rab 11.The images have been analyzed in ImageJ to highlight areas of co-localization. The upper panels are phase, the middle panels are stained with DAPI to show nuclei, and the lower panels illustrate areas where nectin 2 and rab 11 are co-localized (shown in white). Notice that co-localization (pink arrows) is most apparent at stage IX where new adhesion junctions are forming with the next generation of early elongating spermatids. At the same stage, co-localization in basal regions of the epithelium where junctions are beginning to turn over to allow the translocation of spermatocytes from basal to adluminal compartments of the epithelium also is more apparent than at earlier stages. Bar = 50 mm.  	
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    Figure 5.9 Model of junction turnover and recycling in the seminiferous epithelium of the rat based partly on data presented in this chapter. Nectin 2 is internalized into the Sertoli cell at apical tubulobulbar complexes where the internalization machinery is associated with Rab 5. Some of the adhesion protein enters recycling compartments in the cell and is targeted to newly forming adhesion junctions with early spermatids deeper in the epithelium. Rab 11 is associated with nectin 2 containing compartments associated with these early spermatids. We speculate that a similar process may occur at basal junction sites.  	
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    Chapter 6 – Cortactin depletion results in short tubulobulbar complexes and spermiation failure in rat testes. 	
   6.1 Brief synopsis The working hypothesis of this thesis is that tubulobulbar complexes that form between Sertoli cells and spermatids are subcellular machines that internalize intact intercellular adhesion junctions and that the complexes are a key element of the sperm release mechanism. If tubulobulbar complexes are a significant part of the sperm release mechanism, then perturbation of tubulobulbar complex structure/function should lead to spermiation failure. Because there are no culture models of sperm release, the approach taken in this study to verify that prediction is to deplete cortactin, a known component of tubulobulbar complexes in rats, by intratesticular injection of siRNA reagents, determine if the knockdown has an effect on tubulobulbar complexes, and then determine whether there is a spermiation defect. This chapter reports the results of cortactin knockdown experiments. Although tubulobulbar complexes were present in siRNA treated rat testes, they were shorter three days after injection than those in contralateral control testes injected with a scrambled sequence. Moreover, spermiation delay and failure were detected in the knockdown testes, and ectoplasmic specializations (testis specific adhesion junctions) remained associated with those spermatids retained by the  	
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    epithelium. I conclude that apical tubulobulbar complexes are a significant part of the normal process of sperm release and that interfering with their structure/function results in spermiation failure.  6.2 Results 6.2.1 Normal arrangement and structure of tubulobulbar complexes in rat Tubulobulbar complexes that form in apical regions of the seminiferous epithelium are elongate tubular extensions of the attached plasma membranes of the spermatid and Sertoli cell that project into the Sertoli cell in regions previously occupied by unique adhesion junctions known as ectoplasmic specializations. In rats, as many as twenty-four complexes, each ranging from 2-3 µm in length develop in two parallel rows associated with the concave face of the “hook shaped” spermatid heads (Russell and Clermont, 1976a; Russell, 1979d) (Fig. 6.1). The “double-membrane” core of each complex is cuffed by a dendritic actin network (Young et al., 2009a). In addition to actin, other molecular components of tubulobulbar complexes include Arp2/3 (Vaid et al., 2007), N-WASP (Young et al., 2009a), cortactin (Young et al., 2009a), dynamin (Vaid et al., 2007), amphiphysin (Kusumi et al., 2007), and cofilin (Guttman et al., 2004a). Formation of each complex is initiated in the Sertoli cell by a clathrin-coated pit (Young et al., 2009a), which retains an attachment to the neighboring germ cell plasma membrane as the complex elongates and matures (Russell and Clermont, 1976b). In regions of the coated pit where the germ cell plasma membrane 	
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    remains attached to the Sertoli cell, fine filamentous connections occur between the two cells, and the germ cell membrane contains a distinct sub-membrane density (Russell and Clermont, 1976b). As the complex matures a swelling develops in the distal third of the structure just proximal to the coated pit. This bulbar region is not associated with actin. Rather, it is closely associated with a cistern of endoplasmic reticulum. The bulbar region eventually buds from the complex and is internalized by the Sertoli cell (Russell and Clermont, 1976a; Russell, 1979d).  6.2.2 Cortactin knockdown by intratesticular siRNA In the first set of experiments on five animals, cortactin bands on Western blots of whole testis lysates from testes injected with cortactin (Cttn) targeted siRNAs were clearly weaker relative to those of similar lysates from contralateral control testes injected with non-targeting siRNAs (Fig. 6.2). This data made it possible to conclude that cortactin in the testis was successfully knocked down by intratesticular injection of siRNA.  6.2.3 Tubulobulbar complexes are shorter in cortactin knockdown testes Tubulobulbar complexes in sections of stage VII seminiferous epithelium appeared qualitatively shorter in cortactin knockdown testes relative to control testes when double labeled for cortactin (Fig. 6.2 B) and actin.  	
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    The numbers of pixels above a set threshold occupied by the total cluster of tubulobulbar complexes adjacent to the concave face of mature spermatids at Stage VII of spermatogenesis in experimental and control testes labeled for either cortactin or for actin were quantified as an index of tubulobulbar complex length. There was a significant difference between experimental and control values (p < 0.0028 for cortactin and p < 0.0004 for actin) (Fig. 6.2 C,D). To gain a better understanding of the morphological effects of the siRNA, the structures were examined at the ultrastructural level. In electron micrographs of spermatids and adjacent Sertoli cell regions in favorable sections where tubulobulbar complexes could be seen along their entire length, the structures were clearly shorter in experimental (Fig. 6.3 B,C,E,F) than in control tissue (Fig. 6.3 A,D). Moreover, when apical Sertoli processes containing mature spermatids at Stage VII of spermatogenesis were sectioned so that the total cluster of tubulobulbar complexes was cut in cross section in control testes (Fig. 6.4A), many fewer profiles of tubulobulbar complexes were visible in experimental material when cut at the same level (Fig. 6.4 B,C). This is a result consistent with that predicted if tubulobulbar complexes were shorter in experimental than in control testes.  In conclusion, cortactin knockdown resulted in shorter  tubulobulbar complexes.  	
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    6.2.4 Spermatids retained in the epithelium Mature spermatids are released as spermatozoa into the lumen of seminiferous tubules in Stage VIII of spermatogenesis (Leblond and Clermont, 1952). During this spermiation process adhesion junctions (ectoplasmic specializations) between Sertoli cells and spermatids disappear, Sertoli cell cytoplasm is gradually withdrawn from spermatid heads, spermatids detach from the apex of the epithelium, and residual lobes from the released spermatids are retained by Sertoli cells. In light micrographs of the seminiferous epithelium from control testes at Stage VIII, spermatids were being released in the normal fashion (Fig. 6.5 A,B). In the epithelium at stage VIII of experimental animals retraction of the Sertoli cell cytoplasm away from spermatids appeared delayed in some tubules (arrows in 6.5 C). Significantly, after most spermatids in controls had been released (Fig. 6.5 A,B) some spermatids in experimental testes appeared not to be released and were retained by the epithelium (arrow marked by asterisk in Fig. 6.5 C, arrowheads in Fig. 6.6 C,D,E,F). When evaluated ultrastructurally, ectoplasmic specializations were absent from Sertoli cell regions adjacent to spermatids being released from control epithelia (arrowheads in Fig. 6.6 G,H), but were still present in association with spermatids retained at the apex of experimental testes (Fig. 6.6 I,J). In conclusion, there is evidence of a delay and a failure of spermiation in experimental testes, and that adhesion junctions (ectoplasmic specializations) persisted at intercellular attachment sites where spermatids were retained by the epithelium.  	
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    6.2.5 SiRNA reagents enter sertoli cells and result in cortactin knockdown in isolated seminiferous epithelia To verify that the siRNA administered by intratesticular injection actually entered Sertoli cells and resulted in cortactin knockdown in the seminiferous epithelium, additional experiments were run using the same pool of cortactin targeting and non-targeting duplexes but with one nucleotide from each pool modified with DY547 on the 5ʼ end of the sense strand. This nucleic acid tag can be visualized by confocal microscopy using an appropriate laser. The dye was detected in the seminiferous epithelium of all animals that received the modified siRNA and non-targeting reagents (Fig. 6.7 B). To confirm that cortactin was indeed knocked-down in experimental testes, Western blots were run on whole testis lysates from animals using the same reagents as those used for the dye morphological studies. Cortactin bands were weaker in experimental lysates than in contralateral control testis lysates (Fig. 6.7 A). To confirm that cortactin knockdown occurred in the seminiferous epithelium, seminiferous epithelium was isolated from two of the above animals and then analyzed by immunoblotting. In both animals, successful depletion of cortactin was detected (Fig. 6.7 C). As in our previous experiments, the number of positive pixels above threshold was significantly less adjacent to mature spermatids in experimental animals than in controls when sections were labeled for cortactin (Fig. 6.7 D). We conclude that siRNA reagents entered Sertoli cells  	
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    and that the reagents targeting cortactin resulted in lower overall levels of the protein in the seminiferous epithelium than in controls. These results confirm the successful delivery of the siRNA into the seminiferous epithelium.  6.3 Discussion There is a growing body of evidence that tubulobulbar complexes that develop adjacent to mature spermatids are subcellular machines that internalize intact intercellular junctions (Guttman et al., 2004b; Young et al., 2009b; Young et al., 2012). In this study we use intratesticular injection of siRNA reagents to target a known component of tubulobulbar complexes, alter tubulobulbar complex structure and correlate this altered structure with a fundamental biological event during spermatogenesis – sperm release. If tubulobulbar complexes associated with spermatids do in fact internalize junctions, then perturbing their structure/function should prevent junction turnover ultimately delaying or preventing sperm release from the epithelium. In this study an in vivo siRNA approach was used to knockdown cortactin, a protein concentrated at tubulobulbar complexes in the seminiferous epithelium (Young et al., 2009a), and a component of the actin cuff surrounding the central doublemembrane core of each structure. Cortactin is known to coordinate membrane dynamics and cytoskeletal remodeling in other systems – two features that also make it a key player during the process of clathrin-mediated endocytosis (Merrifield et al., 2005; Schafer,  	
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    2002). Two of cortactinʼs five functional domains allow it to simultaneously bind F-actin and Arp2/3. Therefore, cortactin not only promotes nucleation and branching but also acts to stabilize newly formed branches (Weaver et al., 2001). These features, paired with cortactinʼs affinity to bind newly incorporated actin molecules make it a key component in the formation of an actin network (Bryce et al., 2005). Temporarily silencing the expression of cortactin leads to instable actinbased structures in a number of systems. In the absence of cortactin, lamellipodial protrusions retract due to instability (Bryce et al., 2005). Cortactin knockdown in osteoclasts causes a loss of podosome formation (Tehrani et al., 2006). Significantly, podosomes somewhat resemble tubulobulbar complexes in that they are tubular structures with a central membrane core cuffed by a dendritic actin network (Young et al., 2009b). Here we show that when administered to rats by intratesticular injection, siRNA reagents enter Sertoli cells and caused a knockdown of cortactin in the seminiferous epithelium. As observed in other systems where knockdown of cortactin prevented the formation of some structures without altering other cortactin containing features, reduced cortactin levels in the seminiferous epithelium altered the phenotype of tubulobulbar complexes without appearing to effect actin bundles in ectoplasmic specializations where cortactin also has been localized (Kai et al., 2004; Tehrani et al., 2006; Young et al., 2009b).  	
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    When labeled for cortactin and/or actin, tubulobulbar complexes were clearly shorter in epithelia from testes injected with targeting siRNA reagents relative to similar epithelia from contralateral testes injected with non-targeting siRNA reagents. This result was confirmed quantitatively using a threshold method to evaluate the numbers of positive pixels generated by labeling tubulobulbar clusters adjacent to spermatid heads. Using this method, the pixels above threshold remaining in each image were counted and considered an index of tubulobulbar complex length. Numbers of pixels present in tubulobulbar complexes from knockdown animals were significantly different from controls, both for actin and cortactin stained material, supporting the conclusion that cortactin knockdown reduces tubulobulbar complex length. Significantly, in knockdown epithelium a delay in retraction of Sertoli cell away from mature spermatids and spermiation failure was detected. Moreover, in cases where spermatids remained attached to the epithelium, adhesion junctions (ectoplasmic specializations) also were present, an observation consistent with a failure of the normal junction removal mechanism. I propose that short tubulobulbar complexes in the knockdown epithelia cause a failure of the structures to completely internalize intercellular adhesion junctions between Sertoli cells and spermatids and that this results in a delay in and possibly eventual failure of sperm release. Because only one time point (3 days) was evaluated after siRNA injections, it is not possible to distinguish between (1) an initial complete disassembly of tubulobulbar complexes followed  	
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    by partial recovery over the 3 day time period, and (2) a continuous prevention of tubulobulbar complexes to develop to their full length. Collins and coworkers, on the basis of a high resolution ultrastructural study, conclude that one of the actin filament networkʼs primary roles during clathrin-mediated endocytosis is to elongate the bud neck in order to drive the endocytosed vesicles away from the plasma membrane (Collins et al., 2011). This is consistent with the observation of short tubulobulbar complexes after cortactin knockdown. Due to a lack of stability in the actin network caused by a depletion of cortactin, tubulobulbar complexes may not be able to elongate and acquire or maintain normal length and thereby fail to internalize the same amount of junctional membrane. The time period (3 days or roughly 72 hrs) from injection of the siRNA reagents until evaluation of phenotype in this study was sufficient to observe effects on tubulobulbar complexes and on sperm release. In the rat, stage VII of spermatogenesis, the stage when tubulobulbar complexes are most apparent, lasts roughly 56 hours (Russell, 1984). Sperm release generally occurs in late stage VIII and this stage lasts 29 hours. In this study, siRNA reagents effecting stage VII tubules soon after injection would progress through at least to stage VIII when the study was terminated. Also, because tubulobulbar complexes begin development prior to stage VII and because of the length of stage VII, I would anticipate also seeing effects on tubulobulbar complexes in stage VII when the experiment was terminated.  	
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    Interestingly, cortactin homozygous knockout mice appear phenotypically normal and live a normal life span (Schnoor et al., 2011), however, their fertility and testicular morphology have not been reported. Similarly, the anti-androgen flutamide is reported to reduce testicular cortactin levels and to cause phenotypic changes in ectoplasmic specializations (Anahara et al., 2006) but any effects on tubulobulbar complex morphology or on spermiation have not been evaluated. The data presented here are consistent with the hypothesis that tubulobulbar complexes are subcellular machines that internalize intact intercellular adhesion junctions and that their function is essential for normal sperm release. Significantly, these findings introduce a new paradigm for junction internalization generally in cells and link the mechanism to a biologically significant event – sperm release.  	
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    6.4 Chapter 5 figures  Figure 6.1 Position and arrangement of tubulobulbar complexes associated with sertoli/spermatid attachment sites in rat seminiferous epithelium. (A) 1 µm thick section at stage VII of spermatogenesis showing clusters of tubulobulbar complexes adjacent to the concave surfaces of hook-shaped spermatid heads at the apex of the epithelium. Bar = 10 µm. (B) Schematic illustration of tubulobulbar arrangement adjacent to mature spermatids.  	
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    Figure 6.2 Evidence of cortactin knockdown (A) Western blots of seminiferous epithelial lysate from five animals indicate weaker cortactin bands from experimental testes in comparison to control. Calnexin was used as a loading control. (B) Immunofluorescence of stage VII spermatids labeled for cortactin (green) and DAPI (blue). Cortactin staining outlines individual tubulobulbar complexes from experimental and control testes and reveals a difference in length. Bar = 5 µm. (C) Quantitation of knockdown was determined by probing cryosections from control and experimental tissue labeled for cortactin. Mature spermatids were randomly selected and set to a predetermined threshold using ImageJ software. The number of positive pixels 	
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    was determined using the “analyze particles” function. The number of pixels was considered to be an index of tubulobulbar complex length in each cluster. Experimental tubulobulbar complexes were shown to be significantly different from control tubulobulbar complexes (P = 0.0028) (D) Quantitation of knockdown determined by probing cryosections from control and experimental tissue labeled for filamentous actin. Experimental tubulobulbar complexes were shown to be significantly different from those in controls (P = 0.0004).  Figure 6.3 Representative longitudinal sections of tubulobulbar complexes from two different rats. (A, B, C and D, E, F). Relative to tubulobulbar complexes in the control testis of each animal (A, D) similar complexes are noticeably shorter in the cortactin siRNA treated contralateral testes (B, C, E, F). Bar = 200 nm.  	
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    Figure 6.4 Cross sections through apical sertoli cells from control and siRNA treated testes. Cross sections through apical Sertoli cell lobules containing mature spermatids at Stage VII of spermatogenesis in control (A) and cortactin siRNA treated (B, C) testes. The plane of section for each panel is similar to that shown in the right image on Figure 1B. Notice that there are many fewer profiles of proximal tubular regions of tubulobulbar complexes (arrowheads) in the siRNA treated material relative to control tissue. This is because the complexes are shorter in cortactin knockdown tissue relative to controls.  	
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    Figure 6.5 Evidence of delayed spermiation. Light micrographs of Stage VIII seminiferous tubules from control (A, B) and cortactin targeted siRNA (C) injected testes. Notice that in panel (C) Sertoli cell cytoplasm (arrows) still engulfs the spermatid heads (arrowheads) at the apex of the epithelium whereas this cytoplasm has been withdrawn in controls. Also notice that there is one spermatid head (arrow marked with asterisk) that is positioned deep in the epithelium in (C). Bar = 10 µm.  	
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    Figure 6.6 Evidence of spermiation failure and persistence of adhesion junctions in cortactin siRNA testes. (A-F) 1 µM thick plastic sections stained with toluidine blue of stages VIII (late)/IX of spermatogenesis. In sections from control testes (A, B), spermatids are in the final stages of release (arrowhead in (A)) or have been released as spermatozoa from the epithelium (B). In comparable sections from cortactin siRNA testes (CF), numerous spermatids (arrowheads) are observed at the apex of the epithelium, some still surrounded by the apical process of the Sertoli cell (arrowhead in (C)). Others are positioned deeper in the epithelium (arrowheads in D, E, F). Bar = 10 mM. When viewed at the ultrastructural level, spermatids being released from the epithelium at late stage VIII in sections of control testes lack ectoplasmic specializations in Sertoli cell regions adjacent to spermatid heads (arrowheads in (G, H) and apical regions of the Sertoli cell are being withdrawn from the heads. In sections at stages VIII (late)/IX of cortactin siRNA testes (I,J) , ectoplasmic specializations are present adjacent to spermatid heads 	
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    that are positioned in the epithelium similar to those shown in panels D,E and F. Bar = 200 nm.  Figure 6.7 Cortactin knockdown occurs in the seminiferous epithelium and siRNA reagents enter Sertoli cells. (A) Western blots of whole testis lysate from five animals used in the dye studies. Cortactin bands are weaker from experimental testes than from control testes even when the protein concentration is greater as indicated by the calnexin staining. (B) Dye appears as clusters in immunofluorescence micrographs of seminiferous epithelium from testes injected with the modified siRNA and the non-targeting control. Bar = 10 µm. (C) To confirm that cortactin knockdown was in the seminiferous epithelium, two animals used to prepare whole testes lysate were also used to prepare isolated seminiferous epithelium lysate. In both animals, bands from experimental lysates were weaker than from control lysates. (D) The quantitation of cortactin determined by the positive pixels above threshold. The number of pixels was significantly less (P = 0.0032) from experimental testes than from control testes.  	
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    Chapter 7 – Materials and methods  7.1 Animals Animals used in this thesis were reproductively active male Sprague Dawley rats or CD1 mice. They were obtained from Charles River Animal Research Laboratories. The rats were maintained according to guidelines established by the Canadian Council on Animal Care and using protocols approved by the UBC and SFU animal care committees. All experiments were done at least in duplicate using tissue from different animals.  7.2 Reagents The paraformaldehyde was obtained from Fisher Scientific (Ottawa, ON, CAN). The secondary antibodies and phallotoxins conjugated to Alexa fluorochromes were obtained from Invitrogen (Burlington, ON, CAN) and those conjugated  to  horseradish  peroxidase  were  purchased  from  Jackson  ImmunoResearch Laboratories, Inc. (Westgrove, USA). Unless otherwise indicated, all other reagents used in the studies were obtained from SigmaAldrich (Oakville, ON, CAN).  	
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    7.3 Primary antibodies Primary antibodies were obtained from the following companies or laboratories  and  used  at  the  indicated  working  concentrations  for  immunofluorescence, immunoblots and/or electron microscopy: -Rabbit anti-cortactin (Sigma-Aldrich Canada) IF [0.005 mg/ml], WB [0.0005 mg/ml] -Rabbit anti-N-WASP (Santa Cruz Biotechnology) WB [0.0008 mg/ml], IF [0.008mg/ml] -Mouse anti-Clathrin Heavy Chain (Sigma-Aldrich Canada) WB [0.002], IF [0.02mg/ml], EM [0.2mg/ml] -Mouse-anti-vinculin (Sigma Aldrich) WB [0.0004 mg/ml] IF [0.004 mg/ml] -Rabbit- anti-zyxin, B71 (Beckerle Lab) WB [0.0008 mg/ml] IF [0.0025 mg/ml] -Mouse anti-N-Cadherin (Zymed) WB [0.0005 mg/ml], IF [0.005 mg/ml], EM [0.08 mg/ml] -Mouse anti-nectin 2 (Hycult Biotechnology) IF [0.009 mg/ml] -Mouse anti-nectin 3 (Hycult Biotechnology) IF [0.01 mg/ml] -Rabbit anti-α6 integrin (Santa Cruz Biotechnology) IF [0.004 mg/ml] WB [0.0004 mg/ml] -Rabbit anti-EEA1 (Abcam Inc.) IF [0.005 mg/ml] -Rabbit anti-beta actin (Novus Biologicals) EM [0.07 mg/ml] -Rabbit anti-Nectin 2 δ (Takai Lab) IF [0.001 mg/ml] WB [0.0001mg/ml] -Goat polyclonal anti-Rab11 (Santa Cruz) IF [0.005 mg/ml] WB [0.0005 mg/ml]  	
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    -Rabbit polyclonal anti-Rab5 (Abcam) IF [0.001 mg/ml] WB [0.0001 mg/ml] -Rabbit anti-β1-integrin (Millipore) IF [0.002 mg/ml] -Mouse anti-β-actin (Sigma-Aldrich) IF [0.005 mg/ml] -Rabbit anti-Calnexin (Sigma-Aldrich) WB [0.000025 mg/ml]  7.4 Immunofluorescence 	
   7.4.1 Tissue preparation: Isolated testes were removed from anesthetized animals and perfused briefly with warm PBS to clear blood and then perfused with warm fixative (PBS, 3% paraformaldehyde, pH 7.3) for 30 min.  7.4.2 Frozen sections: Fixed testes were frozen onto aluminum stubs with OCT compound (Sakura Finetek USA, Torrence, CA) by immersion into liquid nitrogen. Sections of 5 mm were cut using a cryo-microtome, attached to poly-L-lysine coated glass slides, immediately plunged into -200C acetone for 5 min, air dried, and then processed for immunofluorescence.  	
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    7.4.3 Fragmented material: Fixed testes were decapsulated in PBS and the seminiferous tubules were mechanically separated into small pieces (~1-2 mm cubes) using scalpels and then transferred into a 50 ml Falcon tube. The material was gently aspirated through an 18-gauge needle and then through a 21-gauge needle to fragment the epithelium. Larger material was allowed to sediment for 10 minutes and the supernatant was collected. Using low-speed centrifugation, epithelial fragments were precipitated and then re-suspended in fresh PBS. This material was then added to poly-L-lysine slides. The slides were allowed to incubate in a humidity chamber for five minutes in order for the tissue to adhere to the slide. Excess fluid was removed and the slides were placed in cold acetone (-20 0C) for five minutes and air-dried.  7.4.4 Immunostaining: Slides with attached cryosections were rehydrated and blocked with 5% normal goat serum (NGS) in TPBS-BSA (PBS containing 0.05% Tween-20 and 0.1% bovine serum albumin) for 20 min at room temperature. The primary antibody used to probe cryosections was raised in rabbit, goat or mouse. The primary antibody added to the slides was made up in TPBS-BSA with 1% NGS or NDS (normal donkey serum for studies using antibodies raised in goat), and incubated overnight at 40C in a humidity chamber. The material was washed extensively with the TPBS-BSA (wash buffer), then incubated for 60 minutes at  	
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    370C with secondary antibody conjugated to a fluorochrome (goat antirabbit/mouse AlexaFluor 568 or 488 or donkey anti-goat Alexa Fluor 488 (Invitrogen). The slides were again washed and then coverslips were mounted using Vectashield (Vector Labs, Burlington, ON). The tissue was visualized using a Zeiss Axiophot microscope fitted with appropriate filter sets for detecting fluorescence and with appropriate optics for phase microscopy and an inverted Zeiss Axiovert LSM 5 confocal microscope equipped with epifluorescence and LSM 5 Pascal software. Image acquisition was done with the LSM 5 Pascal software. Filamentous actin was labeled by making up AlexaFluor 568 or 488 phalloidin in TPBS-BSA and staining slides for 20 minutes at room temperature. Slides were then extensively washed in TPBS-BSA.  7.4.5 Antigen retrieval Because the β1 integrin antibodies did not react on tissue processed using standard techniques for immuno-fluorescence, a different fixation protocol was explored and the use of antigen retrieval methods to recover antigenicity. Other studies have used the same antibody to label paraffin sections using antigen retrieval (Beardsley and O'Donnell, 2003). For all β1 integrin experiments, tissue was processed as described above except that the fixative also contained 0.1% glutaraldehyde and slides were not processed through acetone or allowed to air-dry. Instead, slides were immersed in hot (95-100oC)  	
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    sodium citrate buffer (10 mM Sodium Citrate, 0.05%Tween 20, pH 6.0) immediately after letting epithelial fragments settle onto the slides and removing excess fluid. After 5 minutes, the slides were removed from the buffer and immuno-stained. This protocol was also used for some of the Rab5 experiments and found that staining was improved, possibly because the fixative contained a small amount of glutaraldehyde.  7.5 Western blotting Western blots were performed on either whole testis alone, or on whole testis and seminiferous epithelium lysates.  7.5.1 Whole testis lysate For whole testis lysates, testes were removed from rats under deep anesthesia and decapsulated. The whole testis tissue was extensively homogenized in RIPA lysis buffer (150 mM NaCl, 50 mM Tris, pH 7.4, 5 mM EDTA, 1% NonIdet P-40, 1% deoxycholic acid [sodium salt], 10% SDS) with complete, mini – EDTA-free protease inhibitor (Roche, Mississauga, ON, CAN).  7.5.2 Seminiferous epithelial lysate For blots run on both whole testis and seminiferous epithelium, half of each testis was prepared as described above and the other half was cut into  	
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    small pieces in ice-cold PEM/250 buffer (0.2 M Pipes, 0.1M EGTA, 0.1M MgCl2, Sucrose). The small pieces of seminiferous tubules were collected, placed in fresh PEM/250 buffer and epithelium was separated from tubule walls. The isolated sheets of epithelium were collected and pelletted by centrifugation. Pellets were resuspended and extensively homogenized in RIPA buffer. Lysates were loaded into wells of 1-mm-thick 10% SDS-PAGE gels at equal concentrations as determined by protein assay and run according to standard protocols (Laemmli, 1970). Proteins were transferred onto Immobilon-P transfer membrane (Millipore, Billerica, MA) and then washed for 5 minutes at room temperature with TBST (500 mM Tris, pH 7.5, 150 mM NaCl, 0.1% Tween20). The blots were blocked for 12 hours at 40C using 4-5% nonfat milk (Blotto, Santa Cruz Biotechnology, Santa Cruz, CA). Membranes were washed 3 times, 5 minutes each wash, and then incubated for 1 hour at room temperature with primary antibody at the concentration indicated previously (see primary antibodies). Blots were washed 3 times; 5 minutes each wash with TBST and then incubated for 1 hour at room temperature with goat anti-rabbit, goat antimouse or donkey anti-goat horseradish peroxidase-conjugated antibody. Blots were washed 3 times; 5 minutes each with TBST and then reacted with ECL (GE lifesciences, Pittsburgh, PA, USA) and bands were visualized with Bioflex EC film (Interscience, Markham, ON, CAN).  	
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    7.6 Light and standard electron microscopy Testes were removed from anesthetized animals and then perfused, through the spermatic artery, for 2 min with warm (330C) PBS (150 mM NaCl, 5mM KCl, 0.8 mM KH2PO4, 3.2 mM Na2HPO4 pH 7.3) to clear the organ of blood. This was immediately followed by perfusion for 30 min with fixative (0.1 M sodium cacodylate, 1.5% paraformaldehyde, and 1.5% glutaraldehyde, pH 7.3). After perfusion, each testis was cut into small pieces and immersion fixed for an additional 2 hrs. The testis material was then washed with 0.1 M sodium cacodylate for three 10 min washes, then further fixed on ice for 60 min in 1% OsO4 in 0.1 M sodium cacodylate buffer. Following the incubation, the material was washed three times with ddH2O, 10 min each wash, then stained en bloc for one hour with 0.1% uranyl acetate. The material was then washed another three times in ddH2O, then dehydrated in an ascending alcohol series (30%, 50%, 70%, 95%, 100%) for ten minutes at each concentration. This was followed by two incubations of 15 min each in propylene oxide. The blocks then were left in a 1:1 solution of propylene oxide: EM-bed 812 (EM Sciences) overnight. The material was embedded in 100% EM-bed 812 and then incubated at 600C for 24 hours. Thick (1mm) sections were stained with toluidine blue and photographed on a Zeiss Imager.A1 microscope using bright-field optics. Thin sections were viewed and images acquired on either a Philips 300 electron microscope operated at 60 kV or a FEI Tecnai G2 Spirit Transmission electron microscope operated at 120 kV.  	
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    7.7 Electron microscopy Rat testes were removed from animals deeply anesthetized with halothane and perfusion fixed using a two-step fixation protocol and immunolabeling method. Samples were dehydrated through an increasing concentration series of cold (00C to -200C) ethyl alcohols and embedded in Unicryl. Thin sections were cut using an ultramicrotome and collected on nickel formvar/carbon-coated  grids.  To  facilitate  antigen  retrieval,  grids  were  preincubated for five minutes in 6 M urea in 50 mM glycine at pH 7.3 and then washed with TPBS-BSA (Kusumi et al., 2007). Grids were incubated overnight at 40C in primary antibody, washed, and then incubated 1 hour in secondary antibody conjugated to 10-nm colloidal gold diluted 1:25 in TPBS-BSA containing 5% fetal bovine serum. Grids were fixed in 0.1% aqueous uranyl acetate, washed with ddH2O, and then air-dried.  7.8 SiRNA An ON-TARGETplus pool of four siRNAs synthesized by Dharmacon (Lafayette, CO) was used to target rat cortactin (Cttn). An ON-TARGETplus NonTargeting Pool - a pool of four non-targeting siRNAs - was used as a negative control.  	
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    For dye modification studies, the same pool of four oligonucleotides was used with one of the four duplexes having a DY547 modification on the 5ʼ end of the sense strand.  7.9 PEI – mediated siRNA delivery siRNAs were mixed with in-vivo-jetPEI (Polyplus - Illkirch, France) at an N/P ratio of 1/8 at room temperature. For intratesticular injection administration, 50 µl of the Cttn targeting siRNA mixture was injected into right testis and 50 µl of the non-targeting control was injected into the left testis.  	
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    7.10 Animal surgeries  7.10.1 Handling: All animals were handled daily for a minimum of one week before surgical treatment to decrease animal anxiety around investigators and staff. Handling included weighing and a brief physical inspection. A food ʻtreatʼ was given after each handling session.  7.10.2 Surgical preparation: Animals were weighed and anesthesia was induced with 5% isoflurane. Anesthesia was maintained with 2-3% isoflurane for the remainder of the surgical protocol. Surgical sites were shaved and then washed 3 times by surgical soap followed by 70% ethanol. Lacrilube was applied to protect the eyes, and a lactate ringerʼs solution (LRS) was injected subcutaneously to maintain hydration. Pre-emptive pain management consisted of a single dose each of buprenorphine at 0.05 mg/kg and of ketaprophen at 5.0mg/kg injected subcutaneously. In addition, a local analgesia, bupivicaine, was injected subcutaneously along each incision site.  	
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    7.10.3 Surgical procedure: After surgical preparation, animals were transferred to a surgical table equipped with a heat source. Heart rate and oxygen saturation were monitored with a pulse oximeter equipped with a foot clip sensor. Deep anesthesia was ensured by a lack of withdraw reflex upon toe pinch. Using sterile technique, an incision of approximately 1-1.5 cm was made through the right side of the scrotum and skin was opened to expose the spermatic fascia enclosing the testis. An additional incision was made through the fascia to expose the surface of the testis. The testis was positioned to avoid major blood vessels and 32gauge needle attached to a LASI (Hamilton, Reno, NV, USA) syringe system was inserted through the capsule and into the testis. A total volume of 50 µL of Cttn targeting siRNA and PEI mixture was injected into the testis. The needle was withdrawn and the spermatic fascia sutured closed with two to four discontinuous sutures. Subcuticular sutures were used to close the skin incision. Using the same surgical procedure, 50 µL of a non-targeting siRNA and PEI mixture were injected in to the left testis as a control.  7.10.4 Post-surgical care and processing: After surgery, rats were placed in a cage under an infrared heater and allowed to recover. The cage contained clean, dry bedding, hydrogel and bacon softies. In most cases, animals were up and eating within 30 minutes. Animals were then transferred into their home cages with their original cage mates that  	
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    also had the same surgery on the same day. Animals were monitored closely for the remainder of the experiment. After three days, and in each experiment, animals were deeply anesthetized and the testes removed for processing. The testes of two animals were fixed for electron microscopy, those of another two animals were fixed and cryo-sectioned for immunofluorescence analysis and those of a third pair were processed for immunoblotting.  7.10.5 Relative quantitation and statistical analysis of actin and cortactin fluorescence as an indication of tubulobulbar complex length. Quantitation of knockdown was determined by probing cryosections of tissue from control and experimental testis labeled for cortactin or for actin. Ten mature spermatids with associated labeled clusters of tubulobulbar complexes at stage VII of spermatogenesis from each control testis and ten cells from each experimental testis were randomly selected and each cell was set to a predetermined threshold using ImageJ software (Abramoff et al., 2004). When set to a common threshold, the number of positive pixels was determined using the “analyze particles” function. The number of pixels was considered to be an index of tubulobulbar length in each cluster.  	
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    Chapter 8 - concluding chapter In this thesis, I present descriptive and functional data that are consistent with the hypothesis that tubulobulbar complexes in the seminiferous epithelium are responsible for the internalization of intercellular junctions and are a significant component of the sperm release mechanism.  8.1 Overall analysis and conclusions in light of current research  8.1.1 Structure The basic structure of tubulobulbar complexes and the actin-related molecules surrounding them have now been well established. Each complex consists of an elongate plasma membrane core of either a Sertoli cell or of a spermatid that extends into a corresponding tubular invagination of the adjacent Sertoli cell (Russell and Clermont, 1976a). The invagination and the projection of the plasma membranes forms a blind-ended, double membrane tube. An actin network that includes N-WASP, Arp2/3, cortactin and dynamin surrounds the tube (Vaid et al., 2007; Young et al., 2009a). The protein signature of the network suggests that dendritic actin assembly is likely a factor in the formation of tubulobulbar complexes (Young et al., 2009a). A clathrin-coated pit occurs at the tip of the structure and distal parts of the structure expand to form a bulbar region. The bulbar region is not cuffed by the actin network but rather, is closely  	
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    associated with cisternae of smooth endoplasmic reticulum. Also, based on the information about tubulobulbar complexes that has been gathered over the past four decades, a number of observations support the junction internalization theory.  8.1.2 Function The data presented in this thesis demonstrates that the function of tubulobulbar complexes is intercellular junction internalization. In addition to this work, a number of researchers reported data that is consistent with this role. First, tubulobulbar complexes develop at intercellular junctions, indicating that there is a functional relationship between tubulobulbar complexes and junctions (Russell and Clermont, 1976a; Vaid et al., 2007). Significantly, tubulobulbar complexes occur not only at apical junctions between Sertoli cells and spermatids, but also at basal junctions between adjacent Sertoli cells (Russell, 1979c), indicating that the primary function of tubulobulbar complexes likely is not related just to spermatid maturation. Second, junction elements known to be present at ectoplasmic specializations are present in tubulobulbar complexes. Adhesion molecules nectin 2 and nectin 3, known to be present at apical junctions between Sertoli cells and spermatids, also are present in apical tubulobulbar complexes and appear concentrated at their ends (Guttman et al., 2004b). In the rat, α6 integrin (ITGA6) and β1 integrin, adhesion molecules that form a complex at apical sites  	
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    (in the Sertoli cell plasma membrane), also have been shown at the ends of apical tubulobulbar complexes (Young et al., 2009b; Young et al., 2012). At basal tubulobulbar complexes, ultrastructurally identifiable gap and tight junctions have been reported (Du et al., 2013; Pelletier, 1995; Russell, 1979c). In addition, clusters of double-membrane-bound vesicles are observed by electron microscopy at the ends of tubulobulbar complexes (Guttman et al., 2004b) — an observation consistent with the proposal that bulbar regions of the complexes bud from tubulobulbar complexes and “intact” junctions are internalized by Sertoli cells. Third, vesicles associated with tubulobulbar complexes have been shown previously to label positively for lysosomal and endosomal markers including acid phosphatase (ACP2) (Russell, 1979b; Russell, 1979c), lysosomal associated membrane protein 1 (LAMP1) and SGP1 (Guttman et al., 2004b) indicating that tubulobulbar complexes are related in some way to internalization and degradation pathways. In mouse, the endosomal marker EEA1 clearly labels vesicles associated with tubulobulbar complexes (Young et al., 2009a) and Rab 5 appears to label regions proximal to the EEA1 positive regions in rat (Young et al., 2012). Rab 5, an early endosomal marker may be associated with the bulbar region of tubulobulbar complexes because they do not label for actin and are situated precisely at the ends of the strongly actin labeled tubular regions (Young et al., 2012).  Fourth, disruption of tubulobulbar complex formation or failure to form is  	
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    correlated with spermiation delay or failure (D'Souza et al., 2009) (Kusumi et al., 2007). This result would be predicted if tubulobulbar complexes played a role in normal junction disassembly.  8.2 Conclusions based on hypothesis presented in the introduction The data provided in each chapter of this thesis is consistent with the junction internalization hypothesis for tubulobulbar complex function proposed in the introduction. This investigation into the role of tubulobulbar complexes in the seminiferous epithelium begins with a descriptive approach of immunolabeling to gain insight into how they are formed. Current research of homologous structures including podosomes formed by osteoclasts (Ochoa et al., 2000), tubules formed by fibroblasts in a cell-free system (Wu et al., 2010), and the necks of endocytic buds in general (Taylor et al., 2011) indicate that clathrin-coated pits and dendritic actin assembly were likely factors in tubulobulbar complex assembly. Previous work from the Vogl lab localized dendritic actin components Arp2/3 and dynamin 3 (Vaid et al., 2007) to tubulobulbar complexes and the available morphology of tubulobulbar complexes revealed coated pits. I immunologically probed the seminiferous epithelium with two other key components of dendritic actin assembly – cortactin and N-WASP, and with clathrin. All three were present at tubulobulbar complexes leading to the proposal of a model of tubulobulbar  	
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    complex formation that incorporates both the dendritic model of actin assembly and clathrin-mediated endocytosis. Previous work indicating the presence of nectin 2 and nectin 3 in the tubular portion of tubulobulbar complexes (Guttman et al., 2004b) was expanded in this thesis to demonstrate their presence in the vesicles at their ends. Furthermore, these nectin 2 and nectin 3 positive vesicles labeled positively for Early Endosome Antigen 1. An antibody to integral adhesion junction molecule α6 integrin intensely labeled the ends of tubulobulbar complexes and provided more evidence of junction internalization. This chapter also addressed previous reports of the presence of adhesion molecule N-Cadherin at ectoplasmic specializations.  Immunoelectron  microscopy  localized  N-Cadherin  to  desmosome-like junctions in the basal compartment rather than at ectoplasmic specializations. The presence of β1 integrin and early endosome marker Rab 5 at the ends of tubulobulbar complexes further support the junction internalization hypothesis (Young et al., 2012). In immunofluorescence micrographs, Rab5 antibody appears to label regions of tubulobulbar complexes that are lacking actin – a pattern that is consistent with the bulbar region. In addition to finding endosomal markers associated with tubulobulbar complexes in this chapter, evidence of junction molecule recycling is presented. At newly forming junctions deeper in the seminiferous epithelium, nectin 2 colocalizes with recycling marker Rab11 GTPase. This is the first evidence that  	
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    some of the junction protein internalized by tubulobulbar complexes may be recycled to form junctions with the next generation of spermatids. After gathering data to describe the morphology, key components, and possible function of tubulobulbar complexes, enough information about the structures had been obtained to test the prediction that perturbing their formation would result in a delay in sperm release. As mentioned previously, there are no in vitro models that can cultivate mature spermatids and replicate their interaction with Sertoli cells during spermiation. Perturbation of tubulobulbar complexes in vivo has been observed previously on a systemic level by manipulating hormones or knocking out specific genes (D'Souza et al., 2009; Kusumi et al., 2007; McLachlan et al., 2002). The approach that is used in this thesis to perturb tubulobulbar complexes employs a method of intratesticular injection that was first described by Russell (Russell et al., 1987) to assess the toxicity of various agents on testes and to study the mechanisms of normal spermatogenesis. Using this method of intratesticular injection, we took the novel approach of introducing siRNA that is designed to target cortactin, a key component of tubulobulbar complexes. Experimental design included a contralateral control in the non-experimental testis that was a non-targeting, scrambled sequence. Upon knockdown of cortactin, a change in tubulobulbar complex morphology was observed. Tubulobulbar complexes appeared shorter and slightly abnormal in comparison to tubulobulbar complexes from control testes. Quantitation of tubulobulbar complex length revealed a  	
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    significant difference between experimental and control testes. A decrease in the level of cortactin was observed from experimental testis lysate in comparison with control lysate on Western blots. Significantly, spermatids from experimental testes were retained in the epithelium past the stages of normal spermiation – evidence of spermiation failure. No evidence of a delay in sperm release was observed in control testes.  In regards to the proposed function of tubulobulbar complexes serving as a means for removal of cytoplasm, little evidence for this role was observed upon cortactin knockdown. The amount of cytoplasm surrounding spermatids from cortactin-depleted testis appeared to be similar to those from control testis. Also, there did not appear to be a difference in the shape of spermatid heads from cortactin-depleted testis versus those from control testis indicating that the primary role of tubulobulbar complexes may not be related to shaping of the spermatid head.  The successful alteration of the structure of tubulobulbar complexes in this study supported the hypothesis that this altered form was correlated with spermiation failure.  	
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    8.3 Comments on strengths and limitations of the thesis The data provided in this thesis lays fundamental groundwork for understanding the role of tubulobulbar complexes. A major strength of this work is the unique morphological analysis of the seminiferous epithelium that was possible because of methods developed in the Vogl lab. At apical sites, numerous tubulobulbar complexes form in association with each spermatid head and extend into a cuff of Sertoli cell known as a Sertoli cell process. This specialized tissue preparation method involves the fragmentation of fixed seminiferous epithelium that results in breaking the process of Sertoli cell off of the epithelium with tubulobulbar complexes and spermatids remaining intact. When fragmented cells are probed with antibodies of interest, a high-resolution analysis of tubulobulbar complexes is possible. This method was critical to being able to propose a model of formation of tubulobulbar complexes that included both the dendritic model of actin assembly and clathrin-mediated endocytosis. In the past two decades, the field of cell biology has exploded with new discoveries due to the relatively new technology that allows investigators to interfere with the expression of specific genes. It is often possible to transfect cells in vitro with interfering RNA to target genes of interest and look for a corresponding phenotype. Using this type of assay for studying tubulobulbar complexes in the seminiferous epithelium poses a unique challenge.  It is  possible to develop basal tubulobulbar complexes – those formed in the basal compartment at sites of attachment between Sertoli cells – in vitro (Du et al.,  	
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    2013). However, to maintain a primary Sertoli cell culture that can support stage VII-VIII spermatids has not been developed. A recent study used a conditional gene ablation technique to develop cortactin-deficient mice (Schnoor et al., 2011). Although this approach can provide a vast amount of data regarding various systems in the mouse, it is difficult to study tubulobulbar complexes in this animal model. In fact, the fertility of those animals and the phenotype of their testes have not been determined. The ideal model system for studying tubulobulbar complexes is rat because the complexes are best characterized in this model, intercellular junctions are relatively large, and easy visualized in comparison with other mammalian systems. For these reasons, we chose to use an in vivo approach requiring the intratesticular injection of siRNA reagents to achieve a temporary silencing of the expression of cortactin.  8.4 Future directions: As outlined previously, many of the challenges associated with investigating apical tubulobulbar complexes are a result of their unique position in the seminiferous epithelium. When using a gene ablation or knockout approach investigators are confronted with high research costs, various technical challenges, as well as compensatory activation of alternative pathways after  	
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    gene knockdown. In addition, the choice of model system is typically confined to mouse. Organ culture is a method that is well suited to investigating the seminiferous epithelium and can be viewed as a middle ground between in vivo and cell culture methods. Organ culture would maintain the architecture of the seminiferous epithelium and direct it toward normal development. Ideally, germ cell interactions could be observed in situ despite the tissue as a whole being in vitro. In the 1960ʼs and 1970ʼs several research groups used this method but it was abandoned for its inability to induce successful meiosis. Recently, Gohbara et al were able to reevaluate the method using modern equipment and verify complete spermatogenesis (Gohbara et al., 2010). It would be interesting to explore the possibility of organ culture to augment the data from chapter 6 – the effects of cortactin knockdown on tubulobulbar complexes and from chapter 5 – the characterization of vesicles associated with the tips of tubulobulbar complexes and endosomal trafficking of adhesion molecules. An ex vivo system has the potential to take these data a further step and characterize the endosomal pathway by which junctional molecules are trafficked in Sertoli cells. For example, a seminiferous tubule rudiment could be exposed to low temperatures or weak bases to slow endosomal maturation. Two-photon microscopy would allow for imaging of junction molecules tagged with fluorescent markers as they are trafficked through the Sertoli cell.  	
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    In vivo techniques that we have employed have been successful in siRNA knockdown of cortactin in Sertoli cells. Replication of the effect of siRNA knockdown ex vivo, could augment the results. In 2009, Sakai and colleagues performed successful application of siRNA on cultured salivary glands (Gohbara et al., 2010).  A growing number of  investigators are using siRNA successfully in ex vivo experiments. Currently, little is known about the regulation of tubulobulbar complex formation. A recent study found that a number of Sertoli cell miRNAs were upregulated 2-fold after hormone suppression (Nicholls et al., 2011). Not surprisingly, the upregulated miRNAs identified genes that are associated with focal adhesions and with regulation of the actin cytoskeleton. From these data, this group was able to conclude that the effect of hormones acting on Sertoli cells in Stage VIII of spermatogenesis are able to organize and control the expression of miRNAs that regulate cell adhesion and male fertility (Nicholls et al., 2011). These fascinating results open a door to the exploration of how miRNAs work to coordinate the complex timing and events within the seminiferous epithelium and may give insight into how tubulobulbar complexes are regulated. The data presented in this thesis is consistent with the hypothesis that tubulobulbar complexes are subcellular machines that internalize intact junctions during spermiation. Now that this role has been established, it would be interesting to explore other consequences of these massive internalization  	
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    events. For example, what other types of information do Sertoli cells acquire about developing germ cells as these massive adhesion junctions are internalized? Germ cells undergo substantial morphological changes throughout the process of spermatogenesis. Basal and apical tubulobulbar complexes represent two time points where Sertoli cells have an opportunity to obtain vital information from the cells that they are nurturing. A study of spermatogenesis in bovine seminiferous epithelium revealed that vascular endothelial growth factor A (VEGFA) is expressed in both germ cells and in Sertoli cells (Caires et al., 2012). Results from this study demonstrated that treatment of explant seminiferous epithelium with VEGFA stimulated an intracellular response in germ cells that prevents cell death. Sertoli cells appeared to be unaffected by the treatment. It would be interesting to probe tubulobulbar complexes for proteins like caspase 3 or VEGFA that would give Sertoli cells cues about germ cell survival or germ cell death. These cues would possibly indicate, for example, that Sertoli cells might be sampling the cytoplasm of germ cells to acquire information about the population that they are cultivating. This is another of many questions that remain to be determined about the role of tubulobulbar complexes.  	
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    8.5 Discussion of potential applications Interference of the formation of tubulobulbar complexes is linked to spermiation failure, a result consistent with an essential role in sperm release. This information could be vital in understanding various forms of male infertility as well as have potential for contraceptive applications. The delicate interplay of timing, genetics, hormones, mechanical cues and many other factors allow normal, healthy sperm to develop in the seminiferous epithelium. Tubulobulbar complexes represent a small, although highly dynamic part of the process. Despite the challenges in investigating such a complex system, there remain numerous exciting research possibilities and potential applications. However, there is a clear need for more basic science to decipher the process of spermatogenesis before proceeding with clinical application regarding contraception or male infertility.  8.6 Specific contributions A number of specific contributions to various fields of cell biology are represented in this thesis. The identification of actin-based components at tubulobulbar complexes, namely cortactin and N-WASP, led to a proposal of tubulobulbar complex formation. This model is novel in that it incorporates both dendritic actin assembly and clathrin-mediated endocytosis.  	
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    The finding that adhesion molecules at the ends of tubulobulbar complexes associate with endocytic makers and recycling markers introduces the possibility that junction proteins may be recycled to newly forming junctions deeper in the epithelium. A model of junction turnover and recycling was generated based on these results. The main contribution of this work was characterization of tubulobulbar complexes by perturbing their formation in vivo. The short phenotype, persistent ectoplasmic specializations and spermatid retention are consistent with the hypothesis that tubulobulbar complexes are subcellular machines that internalize intact intercellular adhesion junctions and that their function is essential for normal sperm release. Previous to this work, junction turnover in epithelia typically involved disengagement of adhesion molecules and internalization of the junction material into parent cells. Significantly, these findings introduce a new paradigm for junction internalization generally in cells and link the mechanism to a biologically significant event – sperm release.  	
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    Appendix  Appendix: 1 supplemental figures To test the efficiency of the transfection reagent jetPEI in Sertoli cells, we performed an intratesticular injection using jetPEI as the vehicle to transfect Sertoli cells with nectin2-GFP cDNA. Testes were removed after three days and prepared in a similar manner as the immunofluorescence material described previously in materials and methods. Intense green staining at the base of the epithelium was observed indicating that the delivery of cDNA was successful.  Figure A.1 Sertoli cells transfected with nectin2-GFP cDNA.  	
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